plastid protein targeting: preprotein recognition and...
TRANSCRIPT
Plastid Protein Targeting:Preprotein Recognition andTranslocationP. Chotewutmontri*,1,2, K. Holbrook†,1,3, B.D. Bruce*,†,4*Graduate School of Genome Science and Technology, University of Tennessee, Knoxville, TN,United States†University of Tennessee, Knoxville, TN, United States4Corresponding author: e-mail address: [email protected]
Contents
1. Introduction 2282. Origin of Plastids 2293. Plastid Protein Targeting Routes 230
3.1 General Import Pathway 2323.2 Roles of General Import Pathway Components 2343.3 Noncanonical Trafficking 2553.4 Dual Targeting to Plastids and Mitochondria 259
4. Regulation of Plastid Protein Import 2614.1 Organism-Specific Recognition of TPs 2614.2 Expression Control 2694.3 Precursor-Specific Import Pathways 2714.4 Redox Regulation 2724.5 Phosphorylation Regulation 2734.6 Potential Role of Proline Isomerization 2744.7 Regulation by Ubiquitin–Proteasome System 276
5. Concluding Remarks 277Acknowledgments 277References 277
Abstract
Eukaryotic organisms are defined by their endomembrane system and various organ-elles. The membranes that define these organelles require complex protein sorting andmolecular machines that selectively mediate the import of proteins from the cytosol to
1 Contributed equally to the manuscript.2 Current address: Institute of Molecular Biology, University of Oregon, Eugene, OR 97403,
United States.3 Current address: Department of Chemistry and Biochemistry, University of California at Los Angeles,
Los Angeles, CA 90095, United States.
International Review of Cell and Molecular Biology # 2017 Elsevier Inc.ISSN 1937-6448 All rights reserved.http://dx.doi.org/10.1016/bs.ircmb.2016.09.006
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their functional location inside the organelle. The plastid possibly represents the mostcomplex system of protein sorting, requiring many different translocons located in thethree membranes found in this organelle. Despite having a small genome of its own,the vast majority of plastid-localized proteins is nuclear encoded and must beposttranslationally imported from the cytosol. These proteins are encoded as a largermolecular weight precursor that contains a special “zip code,” a targeting sequence spe-cific to the intended final destination of a given protein. The “zip code” is located at theprecursor N-terminus, appropriately called a transit peptide (TP). We aim to provide anoverview of plastid trafficking with a focus on the mechanism and regulation of thegeneral import pathway, which serves as a central import hub for thousands of proteinsthat function in the plastid. We extend comparative analysis of plant proteomes todevelop a better understanding of the evolution of TPs and differential TP recognition.We also review alternate import pathways, including vesicle-mediated trafficking, dualtargeting, and import of signal-anchored and tail-anchored proteins.
1. INTRODUCTION
With rapid advances in genome sequencing, transcriptional profiling,
and proteomics, a comprehensive snapshot of how cells differentially express
their genome is now possible. In plants, systems biology approaches have led
to the generation of a variety of large datasets (Gaudinier and Brady, 2016).
These approaches have revealed the complexity of gene expression and
moving the field forward will require an understanding of the subcellular
localization of the expressed proteins. In plants, these protein dynamics
involve a delicate interplay between not only the nuclear genome and
the genomes of the semiautonomous organelles, the plastids, and the
mitochondria.
Understanding these new challenges comes nearly 40 years after it was
first established that some proteins are targeted to the plastid via a post-
translation mechanism that required a specific N-terminal targeting
sequence (Chua and Schmidt, 1978; Highfield and Ellis, 1978). Despite
the functional similarities, the translocation machineries found in the endo-
plasmic reticulum (ER), mitochondria, and plastids show no similarity in
their core components (Schatz and Dobberstein, 1996). Thus, these import
processes appear to have evolved independently. Surprisingly, in plants cells,
these processes can coexist with little to no missorting. In this review, we
provide a detailed update on our understanding of the structure, function,
and regulation of plastid import pathways, with a focus on the chloroplast
translocons functioning in the two envelope membranes, outer envelope
of chloroplasts (TOC) and inner envelope of chloroplasts (TIC). We also
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look the evolutionary diversity of these molecular machines across a wide
range of phototrophic organisms. We take detailed new look the size and
diversity of the transit peptides (TPs) (Bruce, 2000). Finally, we start to build
a detailed understanding of how TPs can function without sequence homol-
ogy yet perform a common universal function (Bruce, 2001).
2. ORIGIN OF PLASTIDS
Over century ago, Mereschkowsky proposed that plastids derived
from cyanobacteria (Martin and Kowallik, 1999). It is now widely accepted
based on phylogenetic analysis that the primary plastids originated from
endosymbiosis of a cyanobacterial ancestor (McFadden, 2014). Interpreta-
tion of the fossil record and molecular clock analysis date the endosymbiosis
event to over 1.2 billion year ago (Butterfield, 2000; Yoon et al., 2004). The
comparisons of genome organization and sequences of the primary plastids
from Archaeplastida which includes Viridiplantae (green algae and plants),
rhodophytes, and glaucophytes suggest a single endosymbiosis event
(Keeling, 2010; Price et al., 2012; Turner et al., 1999). Aside from the
advantage of photosynthetic capability, the only other endosymbiosis event
resulting in primary plastids was reported in a protist Paulinella (Reyes-Prieto
et al., 2010). To explain the incredibly rare occurrence of primary plastid
endosymbiosis, Ball et al. (2013) showed that the key enzymes required
for the host to utilize photosynthetic carbon were derived from another bac-
terium, the pathogenic Chlamydiales. They further proposed that the endo-
symbiosis occurs only in the infected host where the carbon utilization
becomes beneficial. Many attempts to pinpoint the plastid ancestor suggest
unicellular nitrogen-fixing cyanobacteria based on 16S RNA, photosyn-
thetic, and metabolic gene sequences (Criscuolo and Gribaldo, 2011;
Falcon et al., 2010; Hackenberg et al., 2011; Kern et al., 2011; Pascual
et al., 2011). The most comprehensive study using a large-scale comparison
of 241 complete genomes including 9 cyanobacteria and 4 photosynthetic
eukaryotes indicates that the nitrogen-fixing heterocyst cyanobacteria share
the most genes with the photosynthetic eukaryotes (Deusch et al., 2008).
Although modern heterocyst cyanobacteria, Nostoc sp. PCC7120 and
Anabaena variabilis ATCC29143 harbor around 5500 genes that encode
proteins (Kaneko et al., 2001; Markowitz et al., 2012), plastid genomes of
land plants only encode around 80 proteins (Timmis et al., 2004). This
reduction of plastid genomes was found to occur mainly by the transferring
of plastid DNA to the nuclear genome (Kleine et al., 2009). It was proposed
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that the ability of plastids to import nuclear-encoded proteins enabled the
plastid ancestry to transfer its genes to the nucleus without compromising
its metabolic capacity (Allen, 2003). During the process of endosymbiosis,
the import of proteins encoded by endosymbiont-to-nucleus genes permits
the endosymbiont gene copies to undergo pseudogenization and later loss
(Martin et al., 1993). In Arabidopsis, bioinformatic analysis estimated that
around 4500 protein-encoding genes in the nucleus are originated from
the endosymbiont as shown in Fig. 1 (Martin et al., 2002). Less than half
of these endosymbiont-to-nucleus proteins (about 1800) are targeted to
plastids and the rest functions elsewhere in the cell (Martin et al., 2002).
Interestingly, nonendosymbiont-derived proteins (about 750) also localize
to plastids (Martin et al., 2002) and function in photosynthesis, respiration,
and metabolic pathways (Kleine et al., 2009). While plastid protein import
was crucial in facilitating the endosymbiosis event in the past, it becomes
indispensable to the function of the cells in Arabidopsis where over 2500
proteins in Arabidopsis must gain access to plastids.
3. PLASTID PROTEIN TARGETING ROUTES
Proteins targeted to plastids can be delivered posttranslationally to six
plastid locations: the outer envelope membrane (OM), the intermembrane
Plant cell
Cyanobacterialancestor
Nucleus
Plastid
4500
80
Other locations
1800
2700
750
Fig. 1 Endosymbiont gene transfer and protein targeting to the plastid. Bioinformaticanalysis of Arabidopsis genome sequence (Martin et al., 2002) indicated that around4500 nuclear genes are transferred from the cyanobacterial ancestor and only 87 genesremain in the plastid genome. Although the majority of the proteins encoded by theendosymbiont-to-nucleus genes are targeted to other locations in the cell, about1800 of these proteins are targeted to the plastid. In addition, around 750 proteinsencoded by nonendosymbiont nuclear genes are also targeted to the plastid.
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space (IMS), the inner envelope membrane (IM), the stroma, the thylakoid
membrane, and the thylakoid lumen as shown in Fig. 2 (Keegstra and Cline,
1999; Li and Chiu, 2010). Most of these processes involve protein trans-
location across the plastid membranes: the outer envelope, the inner enve-
lope, and the thylakoid membranes. In general, protein translocation
across or insertion into membrane is mediated by oligomeric membrane
complexes termed translocons (Walter and Lingappa, 1986). The majority
Outermembrane
Intermembranespace
Innermembrane
Thylakoidmembrane
Stroma
Lumen
Direct insertionOEP14 pathwayToc34 pathwayToc159 pathwayOMP pathway
General import
pathway
MGD1/Tic22
Conservative sorting
Stop-trasfer pathway
Spontaneous
Stromalproteins
SRPSec TAT
Noncanonicalpathway
Fig. 2 Plastid protein targeting routes. Proteins targeted to plastids are delivered to sixlocations: outer envelope, intermembrane space, inner envelope, stroma, thylakoidmembrane, and thylakoid lumen. The outer envelope proteins use multiple pathwaysfor targeting while the interior proteins containing TP pass through the envelope(s) viathe General Import Pathway. Some of the proteins contain a second targeting signal forthylakoid lumen targeting. In addition, experimental evidence and proteomic analysisreveal that many proteins utilize noncanonical pathway.
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of plastid-targeted proteins is synthesized as precursors containing the
N-terminal targeting sequences called TPs (Dobberstein et al., 1977;
Kleffmann et al., 2004). Similar to the signal hypothesis (Blobel, 1980),
TPs act as an intrinsic signal on the precursor protein that is recognized
by the targeting receptors associated with the translocons. TPs direct trans-
location of precursor proteins across the double membranes of plastids via
the translocon at the TOC and TIC in a process described as the general
import pathway (Cline and Henry, 1996; Schnell et al., 1997; van’t Hof
and de Kruijff, 1995a). After the precursor is translocated into the stroma,
the TP is readily cleaved allowing the mature domain to fold into its native
conformation or to be further targeted to the thylakoid (Richter and
Lamppa, 2002). So far, the TP has been utilized in targeting precursor
proteins to all of the six locations of plastids and is considered to function in
the general import pathway. This pathway recognizes diverse TP sequences
from various functional groups of proteins. In summary, the general import
pathway functions similar to a central transit hub where the majority of
plastid-targeted proteins pass through before reaching their final locations.
3.1 General Import PathwayThe general import pathway is a working model describing a TP-directed
translocation of proteins into the stroma of plastids (Bruce, 2000). Fig. 3
illustrates the sequential steps in the pathway that reflect the processes that
take place in the cytosol, in both the OM and IM, stromal-localized com-
ponents, and finally the cleavage and processing of the TP. We have not
included steps that are involved in the subsequent targeting to the thylakoid
nor do we address the targeting to the OM, IM, or IMS. Fig. 3 represents a
somewhat selective model that has consolidated many years of work, but
emerging research continues to develop a more complete picture of the
import system.
The OM of plastids has been shown to recruit cytosolic precursors using
multiple pathways. Work performed using isolated chloroplasts shows that
the precursors are able to directly bind to TOC on the surface of chloroplasts
(Fig. 3, step 1a). The binding is reversible when lacking ATP/GTP and den-
oted as energy-independent binding (Jarvis, 2008; Kouranov and Schnell,
1997; Perry and Keegstra, 1994). Addition of GTP also promotes binding
(Inoue and Akita, 2008; Young et al., 1999). TP interaction with chloroplast
lipids suggests that the precursor can directly bind to the lipid surface (Fig. 3,
step 1e) before being transferred to the membrane receptors TOC159 and
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TOC34 (Fig. 3, steps 2f and 3a) (Pilon et al., 1995; Pinnaduwage and Bruce,
1996). Cytosolic factors were shown to interact and recruit precursors to the
membrane receptors. Cytosolic Hsp90 captures precursor (Fig. 3, step 1d)
and delivers it to the membrane receptor TOC64 (Fig. 3, step 2d)
(Qbadou et al., 2006). Phosphorylated TPs interact with 14-3-3 and form
the guidance complex with cytosolic Hsp70 Fig. 3 (step 1c) (May and
90 Å
153015
Precursorprotein
Toc159m Toc34 Toc75 Tic20/21 Tic110 Tic40 Hsp93 cpHsp70
OM
IMS
IM
Stroma
Cytosol
Maturedomain
Transitpeptide
0.1 mM
GTP
SPP
0.1 mM
ATP
4°C
0.1 mM
ATP
25°C
1 mM
ATP
25°C
Binding Translocation
1a
1b
1c
1d
1e
2a
2d
2b
2c
2f3b
3a
4 5 6
78
Toc64 Toc159c Hsp70c14-3-3Hsp90
Fig. 3 The general import pathway. Cytosolic precursors can be targeted to plastidsdirectly by interacting with TOC (1a), or by interacting with the lipids (1e) before trans-ferring tomembrane Toc159 (2f ) and reaching Toc34 (3a). The precursors can also inter-act with cytosolic Toc159 (1b) before transferring to Toc34 (2a). TPs can interact withHsp90 (1d) before being targeted to Toc64 (2d) and further transferred to Toc34(3b). Phosphorylated TPs can interact with the guidance complex (1c) composed of14-3-3 and Hsp70c in cytosol. The precursors from guidance complexes can be trans-ferred to membrane receptors Toc159 (2c) or Toc34 (2b). The binding state of precursorto the chloroplast can be subdivided based on GTP/ATP level and temperature of thesystem (4–5). When the ATP level is greater than 1 mM, the translocation process is ini-tiated by Hsp93/cpHsp70 (6). When the precursors emerge into the stroma, stromalprocessing peptidase (SPP) will cleave TP from precursor proteins releasing the maturedomain (7). TP will be further degraded by PreP1/PreP2 peptidases (8).
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Soll, 2000). The guidance complex was proposed to dock to membrane
receptors TOC159 (Fig. 3, steps 2c and 3a) or TOC34 (Fig. 3, step 2b)
(May and Soll, 2000; Qbadou et al., 2006). Another pathway utilizes the
cytosolic TOC159 pool to deliver precursor to the membrane (Fig. 3, steps
1b and 2a) (Hiltbrunner et al., 2001; Smith et al., 2004).
When low levels of ATP (<0.1 mM) are present, with or without GTP,
the precursor engages in an irreversible binding state, which is referred to as
the early import intermediate (Inoue and Akita, 2008; Kessler et al., 1994;
Kouranov and Schnell, 1997; Olsen and Keegstra, 1992; Perry and Keegstra,
1994; Young et al., 1999). The switching from energy-independent to
energy-dependent bindings seems to involve TOC75 (Paila et al., 2016).
In the presence of 0.1 mM ATP at 4°C, about 110 amino acids are buried
in the translocons (Fig. 3, step 4) (Akita and Inoue, 2009; Inoue and Akita,
2008). When the temperature is increased to 25°C, about 130 amino acids
are buried (Fig. 3, step 5) (Akita and Inoue, 2009; Inoue and Akita, 2008). At
this step, the precursor spans the double membrane via the TOC75 and
TIC20/21 channels and TP interacts with TIC110 (Akita et al., 1997;
Chen and Li, 2007; Inoue and Akita, 2008; Nielsen et al., 1997) forming
the contact site between the double membranes (Schnell and Blobel,
1993). The translocation process is initiated by the ATPase chaperones
Hsp93/cpHsp70 when the ATP level is above 1 mM (Fig. 3, step 6) (Shi
and Theg, 2010; Su and Li, 2010). When the precursor emerges into the
stroma, stromal processing peptidase (SPP) will cleave TP from the precur-
sor, releasing the mature domain (Fig. 3, step 7) (Richter and Lamppa,
1998). TP will be further degraded by presequence peptidases, PreP1/PreP2
(Fig. 3, step 8) (Glaser et al., 2006). The detailed mechanism of processes
such as the specificity of TP recognition and threading of precursors into
the stroma requires further study.
3.2 Roles of General Import Pathway Components3.2.1 TPs and Interacting DomainsTPs are N-terminal extensions that are cleaved during or after import.
They were first identified 40 years ago by Dobberstein and coworkers
(Dobberstein et al., 1977) inChlamydomonas for the small subunit of Rubisco
and named TPs by Chua and Schmidt (1978) to be differentiated from the
signal peptides. TPs are often encoded by the first exon and represent an
addition of new coding information relative to the ancestral gene structure
found in cyanobacteria (Kilian and Kroth, 2004; Tonkin et al., 2008). TPs
are necessary and sufficient to facilitate protein import into plastids; the
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mature domain alone fails to be imported while addition of TP can direct the
import of a nonplastid protein into a plastid (Bruce, 2001). Thus, TP con-
tains information governing the import process. The length of TPs varies
from 20 to 150 amino acids based on the position of the processing site
(Balsera et al., 2009b). However, it has been established that short TPs can-
not direct the import (Bionda et al., 2010). Import only occurs when short
TPs were extended into their mature domains to reach at least 60 amino
acids long (Bionda et al., 2010).
Many attempts have been made to identify the conserved motifs present
amid diverse TPs using primary sequence alignment, but the conservation
becomes greatly reduced with the increasing numbers of TPs analyzed
(Karlin-Neumann and Tobin, 1986; von Heijne et al., 1989). Amino acid
composition and organization are also highly divergent (Bruce, 2000). Nev-
ertheless, three regions were loosely defined: (i) N-terminal domain of about
10 residues, lacking charged amino acids, ending with Pro/Gly and prefer-
ably having Ala as the second residue, (ii) central domain, lacking acidic
amino acids but rich in hydroxylated amino acids, and (iii) C-terminal
domain, rich in Arg and possibly forming an amphiphilic β-strand (Bruce,
2001; von Heijne et al., 1989).
A few studies have determined NMR structures of TPs including those
of TPs of RuBisCO activase (Krimm et al., 1999) and ferredoxin (Lancelin
et al., 1994) from Chlamydomonas, as well as the TP of ferredoxin from a
higher plant Silene latifolia (Wienk et al., 1999). Even though TPs are found
to be unstructured in aqueous solution, they were shown to adopt alpha-
helical structures in membrane-mimetic environments inferring the possible
formation and of an alpha helix in TP recognition (Bruce, 2001). Notably,
the most stable alpha helix of the TP present in higher plant ferredoxin con-
tains a semiconserved FGLK motif that was suggested to interact with the
translocation apparatus (Schleiff et al., 2002; Wienk et al., 2000). This is
an example of the semiconserved physicochemical motifs of TPs that func-
tion as interacting domains determining organelle specificity.
Many groups have determined precursor interactions with translocon
components in intact chloroplast by crosslinking during different stages of
the import process (Akita et al., 1997; Chen and Li, 2007; Inoue and
Akita, 2008; Kouranov and Schnell, 1997; Ma et al., 1996; Perry and
Keegstra, 1994). These interactions can be confirmed in vitro using purified
translocon components and various precursor proteins or TPs. Eleven dif-
ferent precursors (prSSU, prFNR, prOE23, prOE33, prFD, prHsp21,
prLHCP, prRBCA, prE1α, prL11, and prPORA) were used in these
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experiments. These in vitro assays have uncovered many interacting partners
including the guidance complex (May and Soll, 2000), cytosolic Hsp90
(Qbadou et al., 2006), TOC159 (Smith et al., 2004), TOC34 (Schleiff
et al., 2002; Sveshnikova et al., 2000b), TOC75 (Hinnah et al., 2002),
TIC110 (Inaba et al., 2003), stromal Hsp70 (Ivey et al., 2000; Rial et al.,
2000), SPP (Richter and Lamppa, 1999), and presequence peptidases
PreP1/2 (Glaser et al., 2006).
While this interaction information confirmed the cross-linking results
in identifying the combination of translocon components at each state of
the import pathway, it does not provide direct location of the interacting
domains on TPs. Only a limited number of studies reported the interacting
domains of TPs, and these are summarized in Fig. 4. TPs of the small
subunit of RuBisCO (SStp) and ferredoxin (FDtp) are the only TPs with
their interacting domains mapped. The mapped domains include lipids,
TOC159, TOC34, stromal Hsp70 CSS1, and SPP interacting domains
(Becker et al., 2004; Ivey et al., 2000; Lee et al., 2009a; Pilon et al.,
1995; Pinnaduwage and Bruce, 1996; Schleiff et al., 2002; Sveshnikova
et al., 2000b).
At-SStp MASSMLSSAT-MVASP--AQATMVAPFNGLKSSAAFPATRKANNDITSITSNG-GRVNCMNt-SStp MASSVLSSAA-VATRSNVAQANMVAPFTGLKSAASFPVSRKQNLDITSIASNG-GRVQCMPs-SStp MASMISSSAVTTVSRASTVQSAAVAPFGGLKSMTGFPV-KKVNTDITSITSNG-GRVKCMSl-FDtp MASTLSTLSV----------SASLLPKQQPMVASSLPT--NMGQALFGLKAGSRGRVTAM *** : : :. : : * :.:*. : . : .: :.. *** .*
Full length transit peptide
Interacting region
GSRG Pilon et al. (1995)Ps-SPP
MASMISSSAVTTVSRASRGQ Ivey et al. (2000)SAAVAPFGGLKSATGFPV-KKPs-CSS1
MVAPFTGLKSAASFPVSRKQNLDITS *
Becker et al. (2004)Sveshnikova et al.(2000b)Schleiff et al. (2002)
Ps-Toc34
NDITSITSNAt-Toc159
MVAPFTGLKSAASFPVSRKQNLDITSPs-Toc159
VNTDITSITSNG-GRVKCMQV Pinnaduwageet al. (1996)
MASMISSSAVTTVSRASRGQLipid
TLSV----------SASLL Pilon et al. (1995)Lipid
Interacting
partner Reference
Assigned domains in SStp
LipidToc34 Lipid
SPPToc159Hsp70
A
B
C
Becker et al. (2004)
Lee et al. (2009a)
Fig. 4 Interacting domains of transit peptides. (A) Amino acid sequences of full-lengthTPs. SStp, TP of small subunit of Rubisco; FDtp, TP of ferredoxin. (B) Interacting domainsin TPs. For lipid and Ps–CSS1 interaction, the sequences shown in bold have strongerinteractions than sequences with regular characters. Asterisk indicates phosphorylatedSer. (C) All the interacting domains found on SStp. At, A. thaliana; Nt, N. tabacum;Ps, P. sativum; Sl, S. latifolia.
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Mutagenesis (Holbrook et al., 2016; Lee et al., 2002, 2009a; Pilon
et al., 1995), deletion (Holbrook et al., 2016; Kindle, 1998; Kindle and
Lawrence, 1998; Rensink et al., 1998, 2000), Ala scanning (Lee et al.,
2002, 2006, 2008, 2009a), domain swapping (Chotewutmontri and
Bruce, 2015; Chotewutmontri et al., 2012; de Castro Silva Filho et al.,
1996; Lee et al., 2009a; Smeekens et al., 1986), bioinformatics analysis
(Chotewutmontri and Bruce, 2015; Chotewutmontri et al., 2012; Lee
et al., 2008; von Heijne et al., 1989), and the use of synthetic peptides
(Perry et al., 1991; Pinnaduwage and Bruce, 1996; Schnell et al., 1991) have
identified a large number of critical regions in TPs containing information
for the import process. However, the functions of most of these regions are
still unknown. More importantly, these regions seem to be unique
sequences; the same exact sequence could not be found in any two TPs. This
makes it difficult to elucidate their function. These identified regions also
undermine the hypothesis that translocon component recognition of TPs
is based on highly conserved sequence motifs. Instead of exact sequence
motifs, a few import critical regions of TPs can be identified using algorithms
that quantify amino acid composition. These regions are the N-terminal
highly uncharged domain (von Heijne et al., 1989), the Hsp70-interacting
domain (Chotewutmontri and Bruce, 2015; Ivey et al., 2000), and the
FGLK domain (Chotewutmontri et al., 2012; Pilon et al., 1995). Using
quantitative in vitro and in vivo analyses, we established that the location
of a N-terminal Hsp70 interaction domain is required to facilitate interac-
tion with the plastid-localized molecular motors (Chotewutmontri and
Bruce, 2015). This motif is followed by a second region that interacts with
the TOC receptor proteins, which appear to promote TP binding, but can-
not support import alone. This region is termed “FGLK,” which is defined
as having the following: an aromatic residue; a turn-inducing or helix-
breaking residue, a small nonpolar residue, a basic residue, and lacking
any negatively charged residue (Chotewutmontri et al., 2012). Even though
the FGLK motif is semiconserved in TP sequences, its global importance to
the recognition and import of preproteins remains unclear.Many steps in the
general import pathway may recognize TP by physicochemical properties,
which would also explain the lack of conserved motifs in TPs.
3.2.2 Cytosolic and Lipid Modulators of Plastid TargetingPrecursors are proposed to maintain import competency by interacting with
cytosolic chaperones (Jackson-Constan et al., 2001). Multiple studies have
investigated this in bothmitochondria and chloroplasts. One of the common
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proteins identified in both systems are members of the hsp70 class of chap-
erones. In many cases, this work has relied on in vitro studies that show a
direct interaction. Unfortunately, hsp70s are essential cellular proteins and
cannot be deleted or completely downregulated. Although multiple
Hsp70 binding sites have been shown in the majority of TPs (Ivey et al.,
2000; Rial et al., 2000; Zhang and Glaser, 2002) their role both in vitro
and in vivo is not clear. For example, while cytosolic Hsp70 is essential
for the in vitro import of a membrane protein, the precursor of the light
harvesting chlorophyll a/b-binding protein (Waegemann et al., 1990), it
is not necessary for the in vitro import of precursors of soluble proteins such
as ferredoxin (prFD) (Pilon et al., 1990) and the small subunit of RuBisCO
(prSSU) (Dabney-Smith et al., 1999). Clearly more work needs to be done
to elucidate the role of cytocolic hsp70 in plastid protein targeting.
Another biochemically identified cytosolic factor, the guidance complex
composed of 14-3-3 and Hsp70 proteins, improves the efficiency of import
(May and Soll, 2000) 14-3-3 recognizes the phosphorylated TPs (May and
Soll, 2000). This guidance complex was proposed to deliver the precursor
to membrane receptors TOC64, TOC159, or TOC34 (May and Soll,
2000). Later experiments showed that the complex associates with TOC34
but not with TOC64 (Qbadou et al., 2006). Furthermore, some precursors
associate with another cytosolic chaperone, Hsp90. This chaperone delivers
precursors to TOC64 through the interaction between tetratricopeptide
repeats of TOC64 and Hsp90 (Qbadou et al., 2006). The precursor is
subsequently transferred to TOC34 (Qbadou et al., 2006). In addition, TP
receptor TOC159 (Ma et al., 1996; Perry and Keegstra, 1994) has been
suggested to distribute both in soluble cytosolic andmembrane-bound forms
(Hiltbrunner et al., 2001) and it has been proposed that cytosolic TOC159
interacts with precursor proteins and shuttle them to the translocon.
Although cytosolic factors were shown to associate with some precursors
and in certain cases increase import efficiencies, these factors were found to
be nonessential both in organello (Dabney-Smith et al., 1999; Pilon et al.,
1990) and in vivo (Aronsson et al., 2007; Hofmann and Theg, 2005c;
Nakrieko et al., 2004). The rate of in organello protein import without cyto-
solic factors was also suggested to be sufficient to sustain chloroplast devel-
opment (Pilon et al., 1992b). Holbrook et al. (2016) combined detailed
analysis of TP import using in organello and in vivo assays. These analyses
are summarized in Fig. 5 and provide a highly quantitative and resolved bio-
chemical insight while confirming a clear physiological role in living cells.
However, it is clear that there are differential results between the in organello
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and in vivo results. Althoughmutagenesis of a single FGLK physicochemical
domain shows a clear import defect in in vitro assays, mutagenesis of both
domains is required to abrogate in vivo import. This effect is likely due to the
presence of cytosolic factors in the in vivo experiments that can partially
compensate for loss of a TP physicochemical domain.
Fig. 5 In organello and in vivo TP import results. Top (in organello) and bottom (in vivo)heat maps summarize import results using mutants of RuBisCo small subunit TP,removing one or both FGLK physicochemical domains as indicated. Magenta colorcorresponds to a stronger abrogation of import in the tested mutants.
239Plastid Protein Import
ARTICLE IN PRESS
Plastid outer envelope lipids have been implicated to have a role in pre-
cursor targeting and recognition. The composition of lipids making up plas-
tid envelope membranes resembles that of cyanobacteria (Joyard et al.,
1991). These membranes contain high levels of glycerolipids, mono-
galactosyldiacylglycerol (MGDG), digalactosyldiacylglycerol (DGDG),
and sulfolipid (SL), and low levels of phospholipids, phosphatidylcholine
(PC), phosphatidylglycerol (PG), and phosphatidylinositol (PI). The com-
position of the lipids in plastids seems to be maintained over all plastid forms
(Joyard et al., 1991). The presence of galactolipids, MGDG, and DGDG
make the outer membrane of the plastids unique among the cytosolically
exposed membranes (Bruce, 1998).
The roles of lipids in the general import pathway have been analyzed
mainly through the interactions with TPs (Pilon et al., 1995; Pinnaduwage
and Bruce, 1996; van’t Hof and de Kruijff, 1995b; van’t Hof et al.,
1991, 1993). TPs interact specifically with MGDG containing membranes
(Pilon et al., 1995; Pinnaduwage and Bruce, 1996), which has been proposed
to be mediated by hydroxylated amino acids (Pilon et al., 1995). A study
using anArabidopsis dgd1mutant containing less than 10% wild-type DGDG
levels showed that the mutant chloroplasts import proteins to the stroma
slower than the wild-type chloroplasts (Chen and Li, 1998). This import rate
correlated with the reduction of the early import intermediate suggests that
the increased proportion of MGDG in dgd1mutant envelope might trap the
precursor at the energy-independent binding state (Chen and Li, 1998).
Another study using an Arabidopsis MGDG synthase mutant (mgd1-1) pro-
ducing 42% wild-type MGDG levels showed that protein import was not
affected, which may be due to the high levels of remaining MGDG
(Aronsson, 2008). However, how TP–lipid interactions function in the
import process remains unclear. However, the ability of TPs to reorient
MGDG containing bilayers has been hypothesized to be involved (Bruce,
1998; Chupin et al., 1994). Another possibility is based on the observations
that TPs only adopt secondary structures upon binding to lipids. It was
proposed that the TP-chloroplast membrane interaction triggers the folding
of a specific recognition element for protein import (Bruce, 1998).
3.2.3 TOC Apparatus3.2.3.1 Core ComplexThe TOC-TP recognition event is the first step in the general import path-
way in chloroplasts. The TOC core complex is composed of three subunits,
TOC159, TOC75, and TOC34, and forms a heterooligomeric complex
240 P. Chotewutmontri et al.
ARTICLE IN PRESS
ranging from 500 to 1000 kDa (Chen and Li, 2007; Kikuchi et al., 2006;
Schleiff et al., 2003b). The subunits are named based on their predicted
molecular weights in kiloDalton (Schnell et al., 1997). The stoichiometry
of these subunits is still debatable. Ratios of 4:4:1 and 6:6:2 for TOC34:
TOC75:TOC159 have been proposed (Kikuchi et al., 2006; Schleiff
et al., 2003b). This complex size difference also indicates the possibility of
dynamic complex forming higher order structure (Jarvis, 2008). Both
TOC159 and TOC34 contain a GTPase domain (Kessler et al., 1994) while
TOC75 is a beta-barrel protein (Hinnah et al., 1997). The size and compo-
nents of the active TOC core complex are dynamic. Through the course of
plastid differentiation, plastids can increase in size greater than 100-fold and
proteins, lipids, and cofactors necessary for photosynthesis accumulate
(Mullet, 1988). The chloroplast import apparatus must therefore be able
to accommodate the increase in the expression of key photosynthetic pro-
teins. The protein import capability of chloroplasts decreases over the course
of development and has been experimentally determined to decline as much
as 20-fold in in vitro import assays (Dahlin and Cline, 1991). Thus, the core
complex can be modified and its size depends on the metabolic and devel-
opmental state of the cell. Two-dimensional electrophoresis, utilizing
BN-PAGE in the first dimension and SDS-PAGE in the second dimension,
has been shown to be a useful system for the determination of the oligomeric
status of membrane protein complexes (Schagger et al., 1994). We used this
system to investigate the oligomeric states of core complex members
TOC159, TOC75, and TOC34 in developing pea plants. Fig. 6A–F sum-
marizes our analyses. Chloroplasts isolated from young pea plants have more
TOC159 associated with the TOC/TIC supercomplex and more intact
TOC159 participating in complexes in the 1400–630 kDa range (Fig. 6A
and D). The distribution of TOC75 is unchanged throughout the age ranges
(Fig. 6B and D), but young plants have more TOC34 associated with com-
plexes in the 1400–880 kDa range (Fig. 6C and E). This increased associa-
tion of receptors with large complexes suggests that the TOC complexes
may be importing precursors with greater efficiency due to increased bind-
ing. These experiments compare the ratio of TOC components in a partic-
ular size range to the total amount observed, which only allows the
comparison of the proportions of TOC components associated with com-
plexes of differing sizes, but not an overall change in the number of com-
plexes with respect to age. However, it is clear that the core complex is
dynamic and this modulation represents another level of regulation on
the general import pathway.
241Plastid Protein Import
ARTICLE IN PRESS
kDa
200
116.397.4
66.355.4
1320 880 440 132
200
116.397.466.355.4
200116.3
97.466.355.4
7 days old
13 days old
17 days old
1320 880 440 132
200
116.397.466.355.4
kDa
7 days old
200116.3
97.466.355.4 17 days old
200
116.397.466.355.4 13 days old
66.355.4
36.531
21.5
66.355.4
36.531
21.5
1320 880 440 132kDa
66.355.4
36.531
21.5
7 days old
17 days old
13 days old
Super
Comple
x
1400
–630
kDa
Super
Comple
x (86
)
1400
–880
kDa
(86)
880–
630
kDa
(86)
630–
200
kDa
(86)
0
20
40
60 7 days old13 days old17 days old
% T
ota
l To
c1
59
10
30
50
Super
Comple
x
1400
–880
kDa
880–
630
kDa
630–
200
kDa
0
20
40
60
10
30
50
% T
ota
l To
c3
4
Super
Comple
x
1400
–880
kDa
880–
630
kDa
630–
200
kDa
0
20
40
70
10
30
50
60
% T
ota
l To
c7
5
7 days old13 days old17 days old
7 days old13 days old17 days old
A D
B E
C F
Fig. 6 Dynamics of the TOC core complex. (A)–(C) show representative 2D analysis(blue-native electrophoresis and SDS-PAGE) of complexes from chloroplasts isolatedfrom pea plants at 7, 13, and 17 days old. Blots were probed with antibodies againstTOC159 (A), TOC75 (B), or TOC34 (C). (D)–(F) show graphical quantification of the% signal for each complex species at 7, 13, or 17 days old. Quantification was generatedby measuring the pixel intensity of each species using QuantOne software andGraphPad Prism 5.0 was used to generate the graphs.
ARTICLE IN PRESS
3.2.3.2 Identification of TOC Core ComponentsTOC86, the proteolytic form of TOC159, was identified as a major inter-
acting partner of prSSU during binding (Ma et al., 1996; Perry and Keegstra,
1994). Also, antibodies against TOC86 were found to inhibit prSSU bind-
ing to chloroplasts (Hirsch et al., 1994). These findings support a role of
TOC159 as a primary receptor for precursor proteins (Hirsch et al.,
1994; Kessler et al., 1994). Yet, chloroplasts lacking TOC159-binding activ-
ity are able to import prSSU with lower efficiency (Chen et al., 2000). The
intact form of TOC86 and TOC159 was discovered after the Arabidopsis
genome became available (Bolter et al., 1998; Chen et al., 2000).
TOC159 contains three domains, the N-terminal acidic or “A domain,”
the central GTPase or “G domain,” and the C-terminal membrane anchor
or “M domain” (Chen et al., 2000). The A domain is intrinsically disordered
(Richardson et al., 2009) and provides selectivity for precursor protein bind-
ing (Inoue et al., 2010). The G domain is required for insertion of TOC159
into the outer envelope (Bauer et al., 2002; Smith et al., 2002; Wallas et al.,
2003). Recently, the M domain was found to function as a reverse TP.
Unlike TP, which is located at the N-terminus of the proteins, the
M domain is located at the C-terminus of TOC159. It has been shown that
the reverse sequence from C- to N-termini of the M domain fused to the
N-terminal of GFP functioned as TP and was able to direct GFP into the
chloroplast stroma (Lung and Chuong, 2012).
TOC34 was identified from isolated translocons in complex with a pre-
cursor protein (Schnell et al., 1994) and was shown to contain two domains,
the N-terminal GTPase domain (G domain) and the C-terminal membrane
anchor domain (M domain) (Kessler et al., 1994; Koenig et al., 2008a;
Seedorf et al., 1995; Sun et al., 2002). The crystal structures of the
G domain of TOC34 (TOC34G) are related to those of the small GTPases
of the Ras family (Koenig et al., 2008a; Sun et al., 2002). TOC34G also
formed dimers in the crystal where the Arg133 was proposed to function
similarly to the Arg finger of GTPase-activating proteins (GAPs) of Ras
GTPases (Kessler and Schnell, 2002).
TOC75 was discovered at the same time as TOC86 (Perry and Keegstra,
1994). It was shown to be an integral membrane protein (Schnell et al., 1994)
forming 16- or 18 beta-stranded barrel structures (Hinnah et al., 2002; Inoue
and Potter, 2004; Schleiff et al., 2003a; Sveshnikova et al., 2000a). Electro-
physiological measurements show that TOC75 has an intrinsic ability to spe-
cifically recognize TP and has a pore with a diameter at the restriction zone of
about 14 A (Hinnah et al., 2002). The mature TOC75 harbors three repeats
243Plastid Protein Import
ARTICLE IN PRESS
of polypeptide transport-associated (POTRA) domains followed by the
C-terminal beta-barrel domain (Reddick et al., 2008; Sanchez-Pulido
et al., 2003). Although a study showed that the N-terminal POTRA domains
are localized in the cytosol (Sommer et al., 2011), susceptibility of POTRA
domains to proteolysis, and the specific interaction with an IMS component
TIC22 suggest that POTRA domains are in the IMS (Paila et al., 2016).
POTRA domains are shown to have multiple essential roles in TOC75 func-
tion by interacting with TP, TIC22, and the assembly of the TOC complex
(Paila et al., 2016).
3.2.3.3 GTPase Cycle of TOC Receptors: TOC34 and TOC159GTPases exert their control by changing between an inactive GDP-bound
state and an active GTP-bound state, thereby acting as molecular switches
(Gasper et al., 2009). The common denominator of GTPases is the highly
conserved guanine nucleotide-binding (G) domain that is responsible for
binding and hydrolysis of guanine nucleotides. Both TOC34 and
TOC159 belong to the paraseptin group of GTPases, which is most closely
related to the septin GTPase group (Weirich et al., 2008). Similar to other
paraseptin members, TOC GTPases contain the five motifs (referred to as
G1–G5) characteristic for GTP/GDP-binding proteins (Sun et al., 2002).
In addition, paraseptin GTPases contain multiple expansion segments
including additional helices at N- and C-termini, insertions between helix
4 and strand 6 and between strand 6 and helix 5 as shown in Fig. 7 (Sun et al.,
2002; Weirich et al., 2008). The paraseptin and septin GTPase group
together form the septin family which are oligomer-forming GTPases
(Weirich et al., 2008). The dimerization of TOC34 has already been shown
to influence chloroplast protein import (Aronsson et al., 2010; Lee et al.,
2009b). A large number of researchers have focused on elucidating the
GTPase cycles of TOC receptors to further understand the protein import
process, however, some of the results were shown to be conflicting.
TOC34 was crystallized nearly 15 years ago (Sun et al., 2002). Structur-
ally, the overall fold of the TOC34 monomer is divided into a globular
GTP-binding domain (G domain) and a small C-term alpha helix.
TOC34 is remarkably similar in structure to the well-studied small GTPase
p21RAS. Structural superposition of Ras-GppNp and TOC34-GDP
reveals that TOC34 has six sequence inserts I1–I6 (Fig. 7), which except
for I5, have implications in dimerization and GPTase activity (Pai et al.,
1989; Sun et al., 2002). Despite this high-resolution structural data, many
aspects of TOC34 function, as well as the physiological function of GTP
244 P. Chotewutmontri et al.
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---------------------------MTEYKLVVVGAGGVGKSALTIQLIQNHFVDEYDPTIEDSRQAGQLFSLDAAKKKAVESEAEGNEELIFSLNILVLGKAGVGKSATINSILGNQIA-SIDAFGLSTEWVGFQQFPAATQEKLIEFFGKLKQKDMNSMTVLVLGKGGVGKSSTVNSLIGEQVV-RVSPFQAEGEWIGIQQFPPATQSKLLEILGKYKEEDVSSLTVLVMGKGGVGKSSTVNSVIGEKAA-AVSTFQSEGRQIGEMFSLDAAKESASRLEAEGRDDFAFSLNILVLGKTGVGKSATINSIFGETKT-SFSAYGPATEWSGINTFAPATQTKLLELLGNLKQEDVNSLTILVMGKGGVGKSSTVNSIIGERVV-SISPFQSEG
1826
79
79910
HsHRASatToc159atToc33atToc34psToc159psToc34
b1 a1
YRKQ-VVIDGETCLLDILDTAGQEE-------YSAMRDQYMR-----TGEGFLCVFAINN-TKSFETSVREISGTVNGVKITFIDTPGLKSAAMDQSTNAKMLSSVKKVMKKCPPDIVLYVDRLDTQTRDLNLRPVMVSRTMGGFTINIIDTPGLVEAGYVNHQALELIKGFLVNR---TIDVLLYVDRLDVYRVDELLRPTLVSRTRSGFTLNIIDTPGLIEGGYVNDQAINIIKRFLLNM---TIDVLLYVDRLDVYRVDDLTSVTEIVGMVDGVEIRVFDTPGLKSSAFEQSYNRKVLSTVKKLTKKSPPDIVLYVDRLDLQTRDMNPRPVMVSRSRAGFTLNIIDTPGLIEGGYINDMALNIIKSFLLDK---TIDVLLYVDRLDAYRVDNL
408917274
86475
HsHRASatToc159atToc33atToc34psToc159psToc34
b4a2b3b2
G3
DIHQYREQIKRVKDSDDVPMVLVGNKCDLAARTV----------ESRQAQDLA-------------NLPLLRTITASLGTSIWKNAIVTLTHAASAPPDGPSGTPLSYDVFVAQCSHIVQQSIGQAVGDLRLDKQVVIAITQTFGKEIWCKTLLVLTHAQFSPPDE-----LSYETFSSKRSDSLLKTIRAGS-KMRKDRQVVGAITDAFGKEIWKKSALVLTHAQFSPPDG-----LNYNHFVSKRSNALLKVIQTGA-QLKKDLPMLRSVTSALGPTIWRNVIVTLTHAASAPPDGPSGSPLSYDVFVAQRSHIVQQAIGQAVGDLRLDKLVAKAITDSFGKGIWNKAIVALTHAQFSPPDG-----LPYDEFFSKRSEALLQVVRSGA-SLKK
92957135137930138
HsHRASatToc159atToc33atToc34psToc159psToc34
a4b5a3
G4
---RSYGIP--YIETSAKT-R------------QGVEDAFYTLVRE-IRQH--------------MNPSLMNPVSLVENHPLCRKNREGVKVLPNGQTWRSQLLLLCYSLKVLSETNSLLRPQEPLDHQEFEDSAIAVVYAENSGRCSKNDKDEKALPNGEAWIPNLVKAIT-DVATNQRKAIHVDKKMVDGQDLQGFSIPVILVENSGRCHKNESDEKILPCGTSWIPNLFNKIT-EISFNGNKAIHVDKKLVEG-MNPNLMNPVSLVENHPSCRKNRDGQKVLPNGQSWKPLLLLLCYSMKILSEATNISKTQEAADN-DAQASDIPVVLIENSGRCNKNDSDEKVLPNGIAWIPHLVQTITEVA-LNKSESIFVDKNLIDG
1351023195197996198
HsHRASatToc159atToc33atToc34psToc159psToc34
a5b6
G5
I1
I2 I3
I4 I5
I6
180°
Arg130
NN
CC
I6I6
I3 I3
I4 I4
I5I5
HRasHRas atToc33
180°
Arg130
NN
C C
I1I1
I1 I1
I2
I3 I3
I4 I4
I2
I5 I5
atToc33
G1G2
A
B
C
Fig. 7 TOC GTPase domains and Ras alignment. (A) Structural alignment of atToc33(PDB 3BB4) and human p21Ras (HRas; PDB 5B2X) using jFATCAT tool on PDB server.Paraseptin GTPase atToc33 shows expansion segments (in red and cyan) in comparisonto Septin GTPase HRas (in yellow). I1–5, sequence inserts; G1–5, GTP/GDP-binding pro-teinmotifs. (B) Structure of atToc33 as in panel A but without HRas. (C) Alignment of TOCGTPases with HRas. Top label indicates features of TOC GTPases as in panel A. The sec-ondary structure depiction at the bottom was derived from atToc33 (PDB 3BB4),psToc34 (PDB 3BB1), and HRas (PDB 5B2X).
ARTICLE IN PRESS
hydrolysis on protein import, remain unclear. Homodimerization of TOC
receptors are common; atTOC33, psTOC34, and psTOC159 have been
shown to dimerize (Koenig et al., 2008a,b; Oreb et al., 2011; Reddick
et al., 2007; Sun et al., 2002; Yeh et al., 2007). Heterodimerizations between
atTOC33 and atTOC159 and between psTOC34 and psTOC159were also
observed (Bauer et al., 2002; Becker et al., 2004; Hiltbrunner et al., 2001;
Rahim et al., 2009; Smith et al., 2002). Dimerization of atTOC33 was
determined to be involved in the transition of precursor proteins from bind-
ing to translocation state of the import (Lee et al., 2009b).
It is clear that at least three factors have been found to affect dimerization
in vitro. (i) The first factor contributing to dimerization is the protein con-
centration.Monomer–dimerequilibriumof atTOC33andpsTOC34depends
on their concentrations with the dissociation constants (Kd) around 400
and 50 μM, respectively (Koenig et al., 2008a; Oreb et al., 2011; Reddick
et al., 2007). Based on the closed proximity of TOC34 in the core TOC com-
plex (Schleiff et al., 2003b), TOC34 is likely to favor a dimer form in the
complex (Lumme et al., 2014). (ii) Nucleotide loading state is a second factor
that affects dimerization. In both atTOC33 and psTOC34 cases, the GDP-
bound forms produce more dimers than the GTP-bound forms (Koenig
et al., 2008a; Lumme et al., 2014). The same effect was also reported in the
heterodimerization between atTOC159 and atTOC33 (Smith et al., 2002).
Surprisingly, the addition of a GTP transition state analog, aluminum fluoride,
shifted the equilibrium of both atTOC33 and psTOC34 to exclusively
become dimers (Koenig et al., 2008b). These findings and recent work using
FRET interaction analysis indicate that the dimerization is likely triggered by
GTP hydrolysis (Lumme et al., 2014). Lastly, (iii) the effect of TP on TOC
receptor dimerization and function has been the subject of many studies.
GTP-bound forms of atTOC33 (Gutensohn et al., 2000), psTOC34 (Jelic
et al., 2002; Schleiff et al., 2002), and psTOC159 (Becker et al., 2004;
Kouranov and Schnell, 1997) have higher affinities for TP than the GDP-
bound forms. TP is well known to stimulate TOC receptor GTP hydrolysis
(Becker et al., 2004; Jelic et al., 2003; Oreb et al., 2011; Reddick et al.,
2007). Two separate roles of TP in GTP hydrolysis have been reported. First,
it was shown that TP stimulated GTP hydrolysis of psTOC34 while
maintaining the same nucleotide exchange rate suggesting a role of TP as a
GAP but not guanine nucleotide exchange factors (GEF) (Reddick et al.,
2007, 2008). Since GAP and GEF can both increase GTP hydrolysis, but
GAP lowers the transition state energy (Scheffzek et al., 1997)whileGEF stim-
ulates GDP exchange (Bos et al., 2007). Interestingly, another study found that
246 P. Chotewutmontri et al.
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TP stimulated GTP hydrolysis of atTOC33 only when atTOC33 was in
the dimeric form but not in the monomeric form (Oreb et al., 2011). Further
analysis found that TP stimulated atTOC33 GTP hydrolysis through inter-
acting with the dimer and increases the nucleotide exchange rate, which
suggests the role of TP as a GDP-dissociation inhibitor displacement factor,
where each atTOC33 in the dimer acts as GDP-dissociation inhibitor and
TP disrupts the dimer (Oreb et al., 2011). Although the TOC GTPase cycle
has been studied in detail in vitro, less is understood about how the system
works in concert to contribute to precursor import.
3.2.3.4 Models of TOC Receptor FunctionHow TOC receptors function together in protein import is at an early stage
of understanding. There have been several models developed to explain how
these GTPases mediate preprotein recognition and targeting (Bedard and
Jarvis, 2005; Kessler and Schnell, 2004; Reddick, 2010; Smith, 2006).
The “targeting model” (Keegstra and Froehlich, 1999) implicates cytosolic
TOC159 in the capture of precursor proteins in the cytoplasm (step 1b of
Figs. 1–3) and their targeting to the plastid surface (Bauer et al., 2002;
Hiltbrunner et al., 2001; Lee et al., 2003; Smith et al., 2002). TOC159-
precursor complex is proposed to interact with TOC34 (Fig. 3, step 2a)
through the G domains (Bauer et al., 2002). This process is likely controlled
by GTPase cycling, which results in the transfer of precursor to the TOC75
channel to initiate translocation (Smith et al., 2002). In the “motor model”
(Becker et al., 2004), TOC34 on the outer envelope is proposed to act as the
primary receptor (Fig. 3, step 1a). TOC159 in this case is proposed to func-
tion as a GTPase motor pushing precursor proteins through the TOC75
channel (Soll and Schleiff, 2004). The “TOC Clock model” (Reddick,
2010) was developed through analysis of TPs, psTOC159, and psTOC34
interactions together with GTP hydrolysis. It was proposed that in the inac-
tive state, both TOC159 and TOC34 are in GDP-bound forms. Differential
interactions with GTP and TP are hypothesized to modulate the affinity of
the TOC receptors for TP. In this model, TP acts as GAP stimulates TOC34
GTP hydrolysis, and high to low affinity interactions between TOC34 and
TP allows the precursor protein to be imported into the plastid. Recent
studies have classified TOC34 as a member of the GAD GTPase family, a
subclass of small GTPases that are activated by nucleotide-dependent dimer-
ization (Gasper et al., 2009; Lumme et al., 2014). Putative GAD members
have a role in a wide variety of cellular processes, including membrane
fusion and fission (atlastin) (Bian et al., 2011), vesicle trafficking (dynamin)
247Plastid Protein Import
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(Chappie et al., 2010), and cytokinesis (septin) (Sirajuddin et al., 2009).
GADs possess all the structural elements of the classic “switch” mechanism
of small GTPases, but reciprocally activate their catalytic sites in the dimeric
state, rendering classic GEFs and GAPs unnecessary for activity (Gasper
et al., 2009). The proposed GAP function of TPs is not at odds with a
GAD mechanism for TOC34. Based on FRET and PELDOR analysis,
Lumme et al. (2014) suggest that TOC34 can interact with TPs in a dimeric
GDP-bound state, and that GTP alters TOC34 conformation to a “looser”
structure that would allow for precursor protein import through the TOC75
beta-barrel channel. Thus, TP GAP may function as a regulatory switch to
ensure the specificity of plastid import. Ultimately, more detailed analysis is
required to understand how the TOC GTP hydrolysis cycle contributes to
the mechanism of TP recognition and precursor import. Because both
TOC34 and TOC159 hydrolyze GTP, the ability to distinguish the catalytic
events of one TOCGTPase protein over the other would aid in understand-
ing the overall import process. It is likely that the detailed mechanism of
import involves a combination of several of these models, and import reg-
ulation may depend on the metabolic and developmental states of the plant.
3.2.3.5 Preprotein Recognition by TOC GTPasesHow diverse TPs are selectively recognized and processed by the TOC
translocon complex? This is still a paradox, since several 1000 different
TPs exist in a given plant species, yet show very little similarity with each
other—however, they seem to function in a common pathway that involves
the recognition of one or more TOCGTPases, which are proposed to func-
tion as “gatekeepers.” Analysis of plant genomes has revealed that �2100
proteins in Arabidopsis thaliana, and up to �4800 proteins in Oryza sativa
(rice), are imported from the cytosol to their functional location in the plas-
tid (Richly and Leister, 2004). We propose a bimodal model for TP recog-
nition (Fig. 8A), where TPs are recognized at the plastid surface through
interactions between the TOC receptors and the semiconserved physico-
chemical FGLK motif (Holbrook et al., 2016). After GTP hydrolysis, the
TP can access the stroma, where the second recognition event occurs, inter-
action between the N-terminal Hsp70 interaction domain and the stromal
motors (Fig. 8A and B). This two-part interactionmodel would explain how
the TOC translocon is able to manage such a wide variety of TP sequences
(Fig. 8B). Careful analysis of each component of the TOC core complex is
necessary to understand how the complex can balance import specificity
while recognizing a diverse range of TPs.
248 P. Chotewutmontri et al.
ARTICLE IN PRESS
3.2.3.6 Protein Conducting Channel: TOC75Mostmodels of chloroplast protein import suggest that the protein conducing
channel across the outer envelope is a beta-barrel protein that is characteristic
of bacterial outer membranes and the outer membrane of chloroplasts and
mitochondria. The participation of a beta-barrel in protein translocation is
a distinction from the IM, the thylakoid, the ER, and the both membranes
of the mitochondria, which utilize only alpha-helical bundles in their trans-
locons. It has been shown that thepore of theToc complex in the plastid outer
TP model
Stromalinteraction SPP
Spacer
Tocinteraction
HSP70 FGLK
90 Å
15 Å30 Å15 Å
OM
IMS
IM
Stroma
Cytosol
A
B
– – – – ++
++++ ++ ++
or
or
Precursorprotein
Toc159 Toc34 Toc75 Tic20/21Tic110 Hsp93Tic40 Hsp70(CCS1)
Hsp90
PP
GDP
ATP
Hsp70/TP N-terminus
interaction (3)
Chaperone-mediated
molecular motor (4)
TP capture/
Toc translocon unlocking
(2)
TP recognition
(1)
– – – – – –
Fig. 8 Bimodal interaction model for transit peptide recognition. (A) The majority of TPscontains an N-terminal Hsp70-interacting domain (HSP70) linked to a FGLKmotif (FGLK).SPP, the stromal processing peptidase. (B) We proposed the bimodal interaction modelwhere TP is captured by Toc receptor via the FGLK motif. An optimal length spacerallows the stromal molecular motor such as Hsp70 to trap/pull the N-terminal Hsp70-interacting domain to initiate the translocation within a rapid timeframe.
249Plastid Protein Import
ARTICLE IN PRESS
membrane is formed by the TOC75 protein, a member of the OMP85/TPS
(outer membrane protein of 85 kDa/two partner secretion) superfamily
(Andres et al., 2010; Simmerman et al., 2014). This superfamily is found in
outer membranes of gram-negative bacteria, mitochondria, and chloroplasts,
consistentwith their evolutionary origins. TOC75 is comprised of the typical
Omp85/TPS 2-domain architecture, with a N-terminal domain containing
soluble (POTRA) domains and a C-terminal beta-barrel channel formed by
beta strands (Sanchez-Pulido et al., 2003; Simmerman et al., 2014).
TOC75 has been shown to oligomerize with itself and other members of
the TOC core complex (Reddick et al., 2007; Seedorf et al., 1995;
Simmerman et al., 2014). TOC75 is the only member of the TOC core
complex that contains an N-terminal cleavable TP, however, it is a bipartite
sequence (Inoue and Keegstra, 2003). A C-terminal polyglycine stretch
appears to be necessary for retention at the envelope. TheN-terminal region
containing the TP reaches the chloroplast stroma where it is cleaved by sig-
nal processing peptidase. The C-terminal portion of the targeting sequence
spans the IMS and is cleaved by an envelope bound signal peptidase (Inoue
and Keegstra, 2003). Mature TOC75 consists of three soluble N-terminal
POTRA domains and a C-terminal channel comprised of up to 16-beta
strands (Sanchez-Pulido et al., 2003).
In vitro experiments have established thatTOC75 interactswith precursors
during import and is associated with envelope bound import intermediates
(Hinnah et al., 2002; Ma et al., 1996; Perry and Keegstra, 1994; Schnell
et al., 1994). Furthermore, antibodies against TOC75 block protein import
(Tranel and Keegstra, 1996). Patch clamp analysis has established that rec-
onstituted pea TOC75 forms a voltage gated ion channel estimated to be
14 A at its narrowest point (Hinnah et al., 1997). The N-terminal POTRA
domains are proposed to function in TOC complex assembly, precursor rec-
ognition,or chaperoneactivity (Simmermanet al., 2014).Recent findingscon-
firmed the multifunctional roles of POTRA domains in TOC assembly, TP
recognition, and recruitment of TIC22 (Paila et al., 2016).
3.2.3.7 TOC Complex AssemblyThe exact sequence of events in TOC complex assembly remains unclear.
However, studies have established that levels of TOC75 protein are signif-
icantly higher than both TOC34 and TOC159 at early stages of develop-
ment during plastid biogenesis (Kouranov et al., 1998). Therefore, it is
likely that “free” TOC75 is able to nucleate the assembly of newTOC com-
plexes. The TOC75 channel could facilitate interaction between the
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transmembrane helices of the TOC core complex proteins and the lipid
bilayer (Hofmann and Theg, 2005b; Richardson et al., 2014). Analysis of
atTOC75 revealed that POTRA domains recruit atTOC33 and atTOC159
and the deletions of POTRAdomains altered TOC complex assembly (Paila
et al., 2016), supporting the nucleation role of TOC75. Reconstitution of
the complex from individual components has established that TOC159
insertion is dependent on both TOC75 and TOC34, suggesting that
TOC159 is the final element of the core complex assembly (Wallas et al.,
2003). Interactions between the G domains of the TOC34 and TOC159
receptors likely maintain the stability and stoichiometry of the complex.
This assembly mechanism would suggest that the receptor integration deter-
mines the rate of formation of new TOC complexes (Richardson et al.,
2014). However, analysis of TOC complexes suggests that core complexes
can be structurally distinct based on developmental stage and metabolic con-
ditions (Fig. 6A–F). However, TOC75 is the common element in TOC
complexes. Although the measured size of TOC complexes are heteroge-
neous, most complexes are found at over�800 kDa, suggesting that the core
complex contains two molecules of TOC159 and six molecules each of
TOC34 and TOC75 (Chen and Li, 2007; Kikuchi et al., 2006; Schleiff
et al., 2003b) (Fig. 6A–F). Excess TOC75 could provide a pool of nucleating
protein for rapid assembly of distinct complexes, with varying integration of
TOC34/TOC159 family members. Additional analysis of the targeting of
the individual TOC core components plastid targeting and expression is
required to fully understand the factors that control complex assembly.
3.2.4 TIC ApparatusThe TIC complex was identified to be composed of TIC20, TIC21, TIC22,
TIC32, TIC40, TIC55, TIC62, and TIC110 (Bedard and Jarvis, 2005;
Kessler and Schnell, 2006; Smith, 2006; Soll and Schleiff, 2004). Their func-
tions in protein import range from facilitating precursor translocation
through IMS, TOC–TIC formation, TIC channel formation, stromal
motor function, and regulation of import. Many TIC subunits have been
hypothesized to form the translocation channel of the inner envelope. How-
ever, the first proposed TIC channel was identified via electrophysiological
measurement, called protein import-related anion channel (PIRAC) (van
den Wijngaard and Vredenberg, 1999). PIRAC opening is regulated by
TP (Dabney-Smith et al., 1999; van den Wijngaard et al., 1999) and it
was found to associate with TIC110 (van den Wijngaard and
Vredenberg, 1999). The other electrophysiological experiment identified
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TP regulated cation channel TIC110 as a TIC channel (Heins et al., 2002).
This result excludes PIRAC as functioning directly as the TIC channel since
all other protein translocation channels are cation selective (Heins et al.,
2002). It was later found that TIC110 contains six transmembrane helices
and the opening was regulated by Ca2+ (Balsera et al., 2009a). TIC110 is
also essential for TIC complex formation (Inaba et al., 2005). The other
potential TIC channels are TIC20 and TIC21, which are related to the
pore-forming subunits of the mitochondria translocon in the inner envelope
(Reumann and Keegstra, 1999; Teng et al., 2006). The attic20 RNAi
knockdown and attic21 knockout plants showed defective translocation
across inner membranes (Chen et al., 2002; Teng et al., 2006). While
TIC20 is expressed mainly in early development, TIC21 is expressed at a
later stage indicating a shared function of these proteins (Teng et al.,
2006). Components of the TIC system still require additional research to
fully understand their role in the import cycle.
3.2.5 Stromal MotorsIt is well established that internal ATP hydrolysis is the driving force for
plastid protein translocation (Theg et al., 1989). Since the identification
of Hsp70 family of proteins involved in protein import into ER and mito-
chondria (Chirico et al., 1988; Deshaies et al., 1988; Murakami et al., 1988;
Zimmermann et al., 1988), ATPase chaperones have been proposed to drive
the translocation of precursor proteins into plastids (Keegstra and Cline,
1999; Marshall et al., 1990). Two classes of chaperones, Hsp70 and Hsp100,
have now been established to associate and facilitate the protein translocation.
ThreeHsp70proteins associatedwith chloroplastswere first identified frompea
OMand stroma (Marshall et al., 1990). Later, aHsp70was identified in the pea
chloroplast OM as an import intermediate-associated protein (IAP) (Schnell
et al., 1994). This IAP can be detected by antibodies against mammalian
cytosolic Hsp70, mAb SPA-820 (Schnell et al., 1994). Because the Hsp70
IAPwas protected fromanOMimpermeable protease thermolysin, itwas pro-
posed to localize in the IMS and interact with incoming precursor proteins
translocated through TOC75 (Schnell et al., 1994). The gene corresponding
to the Hsp70 IAP has not yet been identified. A later study, however, found
that the protein recognized by mAb SPA-820 is located in the stroma
(Ratnayake et al., 2008). The other outer membrane Hsp70s from pea and
spinach, the thermolysin-sensitiveCom70s, were shown to associatewith var-
ious precursors (Ko et al., 1992; Kourtz andKo, 1997). Com70 also has higher
sequence similarity with mammalian cognate Hsp70s than Escherichia coli
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Hsp70, DnaK (Ko et al., 1992). It was later suggested that Com70 is the
cytosolicHsp70-1 (GuyandLi, 1998;Li et al., 1994).A stromalHsp70was later
shown to associate with the translocation complexes (Nielsen et al., 1997).
In Arabidopsis, the only two Hsp70s predicted to harbor TP were shown
to localize in the stroma (Ratnayake et al., 2008; Su and Li, 2008; Sung et al.,
2001). These stromal Hsp70s are closely related to the bacterial Hsp70,
DnaK (Ratnayake et al., 2008) and were named atcpHsc70-1 and
atcpHsc70-2 (Su and Li, 2008). These cpHsc70s were shown to be directly
involved in protein import. The atcphsc70-1 atcphsc70-2 double knockout is
lethal and the single knockout plants showed reduced protein import effi-
ciencies (Su and Li, 2010). Biochemical analysis also found atcpHsc70s stably
associate with the translocons and the precursors (Su and Li, 2010). The stro-
mal Hsp70, ppHsp70-2 involved in protein import was concurrently iden-
tified in moss Physcomitrella patens (Shi and Theg, 2010).
The stromal Hsp93 (ClpC), a member of the Hsp100 chaperone family,
was first identified to be stably associated with the translocons (Nielsen et al.,
1997). Two homologs were found in Arabidopsis, atHsp93-V and atHsp93-
III (Kovacheva et al., 2005). Crosslinking data suggested that Hsp93,
TIC110, and TIC40 function in concert during protein import (Chou
et al., 2003), consistent with genetic analysis results (Kovacheva et al.,
2005). The molecular interactions between these proteins have been stud-
ied. The binding of TP to TIC110 triggers the association of TIC110 with
TIC40, which in turn induces the release of TP from TIC110 (Chou et al.,
2006). TIC40was also shown to stimulate Hsp93 ATP hydrolysis. Because it
has lower affinity to ADP-bound Hsp93, TIC40 was suggested to act as an
ATPase-activating protein of Hsp93 (Chou et al., 2006). Nonetheless, a
mutant of Hsp93 that obviates its ability to interact with Clp proteolytic core
demonstrates the role of Hsp93 in quality control (Flores-P�erez et al., 2016).BothHsp70 andHsp100 systems are essential for viability of plants (Shi and
Theg, 2011). Knockout of each system are lethal such as those observed in
atcphsc70-1 atcphsc70-2 double knockout (Su and Li, 2008), athsp93-V
athsp93-III-2double knockout (Kovacheva et al., 2007), and pphsp70-2knock-
out (Shi and Theg, 2010). However, these results may reflect not only their
roles in protein import but also other roles in plant development (Constan
et al., 2004; Lee et al., 2007; Su and Li, 2008). In minimally invasive cases,
the knockout lines of a single homolog of each system, such as atcphsp70-1,
atcphsp70-2, and athsp93-V single mutants, import efficiencies dropped to
about 40–60% (Su and Li, 2010) indicating important roles in protein import
of each system.When the functions of both systemswere reduced byknocking
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out a single homolog from each system such as the atcphsp70-1 athsp93-V dou-
ble knockout plant, import was further reduced to 30% (Su and Li, 2010). This
additional reduction compared to the effect of each single mutant, together
with the coimmunoprecipitation ofHsp70 andHsp93 suggested that both sys-
tems act in concert in the same complex (Shi and Theg, 2011).
Interesting, quantitative analysis to determine the energetics of preprotein
import have shown it to be an energy intensive process. Using two different
precursors prepared by three distinct techniques, Shi and Theg have shown
that the import of a precursor protein into chloroplasts is accompanied by
the hydrolysis of �650 ATP molecules. This translates to a ΔG protein
transport of some 27,300 kJ/mol protein imported. When the cost of
preprotein import is expanded to include the complete biogenesis of the
organelle, they estimate that protein import across the plastid envelope
membranes consumes �0.6% of the total light-saturated energy output of
the organelle (Shi and Theg, 2013b).
3.2.6 PeptidasesThe processing of prSSU was discovered nearly 40 years ago (Dobberstein
et al., 1977). This finding led to the identification of two SPPs from pea
(Oblong and Lamppa, 1992; VanderVere et al., 1995) and one in Arabidopsis
(Richter and Lamppa, 1998; Zhong et al., 2003). SPP is classified as a mem-
ber of the metalloprotease M16B subfamily with a zinc-binding motif
(Aleshin et al., 2009; Richter and Lamppa, 1998; Richter et al., 2005;
VanderVere et al., 1995). It was shown that SPP is the general processing
enzyme of plastid protein imports by its ability to proteolyze various precur-
sors (Richter and Lamppa, 1998). SPP specifically binds to the C-terminal
12 residues of TP of prFD (Richter and Lamppa, 1999, 2002, 2003). In
vitro, the TP directs precursor binding to SPP (Richter and Lamppa,
1999). The mature domain is released immediately after cleavage of TP
while TP remains attached (Richter and Lamppa, 1999). TP is further frag-
mented before release from SPP (Richter and Lamppa, 1999). The SPP
recognition sites at theC-terminus of TP seem to share aweak-bindingmotif
(Emanuelsson et al., 1999; Gavel and von Heijne, 1990). Later, the physico-
chemical properties at specific residues were proposed to form a SPP-binding
motif (Richter and Lamppa, 2002). In fact, the regions at the C-termini of
TPs show a positive net charge and are conserved at the position �1 for
basic residue and positions�4,�3,�2, and+1 for uncharged residues related
to the SPP cleavage sites (Richter and Lamppa, 2002).
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Free cleaved TPs in the stroma are potentially harmful (Glaser et al.,
2006) and are degraded by presequence peptidases (PrePs) (Bhushan
et al., 2005). PrePs were originally identified from the mitochondrial matrix
as mitochondrial presequence degradation enzymes (Stahl et al., 2002).
They are classified as members of the metalloprotease M16C subfamily
(Glaser et al., 2006). Arabidopsis has two PrePs, atPreP1 and atPreP2. Both
proteins were found to have dual localizations in both chloroplasts andmito-
chondria (Bhushan et al., 2003, 2005;Moberg et al., 2003; Stahl et al., 2002).
In addition, these proteins have different tissue-specific expression patterns;
atPreP1 is expressed highly in flowers and can be detected in siliques while
atPreP2 is expressed in leaves, shoots, and roots (Bhushan et al., 2005).
3.3 Noncanonical TraffickingAlthough the general import pathway directs the transport of thousands of
plastid-targeted proteins, noncanonical pathways are critical for organelle
function. The chloroplast proteomic data suggest that around 20% of chlo-
roplast proteins lack any predictable targeting sequence (Kleffmann et al.,
2004) and around 1–8% of chloroplast-targeted precursors contain an
ER signal peptide (Kleffmann et al., 2004; Zybailov et al., 2008). Thus,
a small fraction of protein lacking TPs are still able to target to the plastids.
In recent years, several alternate targeting pathways have been reported
(Fig. 9). Noncanonical transport pathways include vesicle-mediated traf-
ficking, import of tail-anchored (TA)/signal-anchored (SA) membrane
proteins, and import of proteins with dual targeting signals to the mito-
chondria and plastid. In this section, we summarize the current understand-
ing of noncanonical transport, although many aspects of these pathways
remain unclear and the focus of ongoing research. Developing a clearer
picture of the integration of noncanonical transport with the general
import pathway will provide insight into plastid biogenesis and organelle
maintenance.
3.3.1 Vesicle-Mediated TraffickingOrganelles cannot synthesize every molecule they require, so a variety of
metabolites, lipids, and proteins must be exchanged via the secretory pathway,
which iswell studiedbetween theER,Golgi, andplasmamembrane.Accumu-
lating evidence suggests that both a cytosol to plastid and intraplastid vesicle
delivery system exist for both lipids and proteins (Karim and Aronsson,
2014; Khan et al., 2013; Lu, 2016). In agreement,Arabidopsis genome analysis,
bioinformatics, and web-based localization prediction tools have confirmed
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that there are plastid-localized proteins with high sequence similarity to the
components of the cytosolic vesicular system (Khan et al., 2013). Analyses have
identified eight COPII-related proteins, as well as proteins related to SNAREs
and RABs (Andersson and Sandelius, 2004; Brandizzi, 2011). Furthermore,
active GTPase homologs to SAR1 and dynamin were identified (Khan
et al., 2013). Thus, intraplastid vesicular trafficking appears to use a system anal-
ogous toCOPII-mediated transport.However, even though putative compo-
nents of the intraplastid vesicular trafficking mechanism have been identified,
substantial research is required to understand their specific roles in transport.
The evolutionary basis for plastid vesicle trafficking has been the focus of
several studies. Interestingly, several of the proteins implicated in plastid vesic-
ular trafficking are conserved in cyanobacteria (Keller and Schneider, 2013;
Cytosol
(2) SA/TA targeting
(1) Vesicle-mediated
transport
(3) Dual targeting
ER/Endomembrane system
Mitochondria
General import pathway
Chloroplast
Vesicle Sec61complex
Ribosome mRNA OM protein
dTPprotein
TOC/TIC
TOM/TIM
Cytosolic factors
Vesiculartransport factors
Thylakoid/IMS protein
Fig. 9 Noncanonical trafficking pathways. Schematic depicts noncanonical plastid traf-ficking pathways: (1) vesicle trafficking of a protein from the ER/endomembrane systemto the plastid; vesicle trafficking of a protein/lipids from the IM to the thylakoid; (2) SA/TA protein cytosolic translation and insertion into the plastid OM; and (3) dual targetingof a protein to the mitochondria and plastid through the TOM/TIM (mitochondria) andTOC/TIC complexes.
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Khan et al., 2013), implying that vesicle trafficking may have developed from
endosymbiosis. However, there are also clear similarities with the cytosolic
system, suggesting possible transfer of secretory pathway components to the
plastid via divergent evolution (Vothknecht and Soll, 2005). Analysis of organ-
isms from diverse lineages suggests a late evolutionary development of plastid
vesicular trafficking (Westphal et al., 2003). However, new phylogenetic
analyses of A. thaliana and O. sativa suggests that vesicular trafficked proteins
are of eukaryotic (not cyanobacterial) origin (Gagat et al., 2013). The early
eukaryotic host-derived proteins were already targeted to the endomembrane
system by their signal peptide and preadapted to be targeted to plastids via the
same system. Furthermore, endomembrane transport was the only viable plas-
tid import route for several eukaryotic proteins that require posttranslational
modifications or dual targeting to the plastid and cell wall. The phylogenetic
results are consistent with a later development of endomembrane trafficking
to plastids (Gagat et al., 2013). It is likely that the system developed in response
to selective pressure to maintain a complex thylakoid membrane (Karim and
Aronsson, 2014), which is spatially separated from the chloroplast envelope
by the aqueous stroma in higher plants.
The lipids that comprise the thylakoid are synthesized through a pathway
that involves exchange of lipid precursors from the ER to the IM; the final
lipid products are produced in the IM but must reach their destination
in the thylakoid (Douce and Joyard, 1990). Although there are several
hypotheses for lipid transfer, both ultrastructural and biochemical evidence
supports a vesicular-mediated trafficking system for lipid movement to the
thylakoid (Karim and Aronsson, 2014). Although vesicular lipid movement
has been characterized, protein targeting through vesicles is less defined.
Furthermore, it remains unclear if vesicles only play a role in intraplastid
localization of proteins, or if proteins can be targeted from the cytosol to
the plastid via vesicles. Two such proteins that utilize vesicle trafficking from
the cytosol contain signal peptide for ER targeting, carbonic anhydrase 1
(Villarejo et al., 2005), and nucleotide pyrophosphatase/phosphordiesterase
1 (Nanjo et al., 2006), were shown to localize to plastids. Additionally, a
recent study searched for nuclear-encoded proteins that may be trafficked
to the thylakoid via vesicles. Bioinformatic analysis identified homologs
to thylakoid-localized proteins with a COPII selection motif and found
21 transmembrane and 12 soluble candidates (Khan et al., 2013). About
45% of these candidates are linked to photosynthesis, implying that a subset
of nuclear-encoded proteins are trafficked to the thylakoid via vesicles (Khan
et al., 2013). Furthermore, RNAi knockdown of a candidate SAR1
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homolog GTPase (cspar1) resulted in downregulation of several nuclear-
encoded photosynthetic proteins, suggesting they are potentially trafficked
via vesicles (Bang et al., 2012). Thus, both bioinformatics and experimental
data support a role for vesicles in both intraplastid delivery and cytosol to
plastid-mediated protein transport. Developing a better understanding of
this system could have important implications in plastid biogenesis, thyla-
koid maintenance, and assembly of the photosynthetic apparatus (Lu, 2016).
3.3.2 SA and TA ProteinsOuter membrane proteins play a key role in intracellular communication,
including exchange of metabolites and lipid synthesis. At least six pathways
mediate protein targeting to the outer envelope of plastids (Hofmann and
Theg, 2005a; Jarvis, 2008;Richardson et al., 2014) (Fig. 2). Furthermore, cor-
rect localizationof plastidoutermembraneproteins is critical for the functionof
the general import pathway. Outer membrane proteins are a group of diverse
proteins that can be divided into alpha-helical transmembrane domain (TMD)
containing proteins and beta barrels consisting ofmultiple transmembrane beta
strands (Inoue, 2015). TMD containing proteins can be further classified based
on the number and location(s) of their TMDsequence (N-terminal,middle, or
C-terminal) (Lee et al., 2014). These proteins are nuclear encoded and synthe-
sized on cytosolic ribosomes; they have distinct, specific requirements for post-
translationally targeting to their functional location. In plastids both SA andTA
proteins play an important role in outermembrane communication. Although
bothSAandTAproteinshavehydrophobicTMDdomains thatdirect insertion
into the plastidOM, the secondary signals thatmediate organelle specificity are
not well understood. Physicochemical domains near the TMD, such as posi-
tively chargedresidues, seemtoplay a role in trafficking (Leeet al., 2014).Cyto-
solic factors likely play a role in the posttranslational insertion of SA/TA
proteins. SomeOMproteins also contain a specializedTP sequence.For exam-
ple,TOC75utilizes a bipartite targeting sequence composedof stromalTPand
envelope targeting signal (Inoue and Keegstra, 2003; Tranel and Keegstra,
1996). The insertions of OEP14, TOC159, and TOC75 also require some
components of the general import pathway. Other outer envelope targeting
pathways have been shown but are not well characterized (Hofmann and
Theg, 2005a).
The most studied pathway for OM proteins requires an N-terminal
TMD targeting signal anchor, which is found in proteins such as the outer
envelope protein of 14 kDa (OEP14) and the TOC subunit of 64 kDa
(TOC64) (Hofmann and Theg, 2005b; Lee et al., 2001). SA proteins are
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a class of OEPs with a single TMD that acts as both the localization signal and
membrane anchor (Rapoport, 2007). SA proteins lack a cleavable TP; a
moderately hydrophobic TMD acts as the primary targeting signal. Another
required motif for targeting of SA proteins appears to be a C-terminal pos-
itively charged domain (Rapaport, 2003). However, the distinguishing the
similar localization signals for plastids vs mitochondria has proven challeng-
ing (Lee et al., 2011, 2014). Another pathway involves a C-terminal “tail
anchor” TMD targeting sequence such as those present in TOC34 and
TOC159 (Smith et al., 2002; Tsai et al., 1999). TA proteins contain a single
C-terminal TMD and basic residues in a C-terminal sequence. Similar to SA
proteins, TA proteins lack a cleavable TP. Plastid TA sequences do not share
any obvious conserved sequence motifs, but they have significantly higher
hydrophobicity index vs mitochondrial TA proteins (Lee et al., 2014).
Only a few plastid TA proteins have been studied in depth, including
TOC complex receptors TOC33 and TOC34. The C-terminal TMD
domain of these proteins is necessary but not sufficient for OM localization;
the GTPase domain of these proteins is required as well (Dhanoa et al.,
2010). However, the exact targeting mechanism is unclear, because a trun-
cated form of the TOC159 GTPase domain lacking a TA domain still cor-
rectly localizes to the OM (Smith et al., 2002). Thus it appears that the TA
sequence is not always sufficient and other sequence context is necessary for
correct localization. The mechanisms that regulate SA/TA protein organelle
specificity and insertion will be the key to understanding organelle biogen-
esis and plant development in depth.
3.4 Dual Targeting to Plastids and MitochondriaAnalogous to plastids, mitochondria import more than 1000 proteins that are
nuclear encoded and synthesized in the cytosol as precursor proteins
(Murcha et al., 2014). The mitochondrial protein import complex is a set
of multisubunit protein complexes that recognize and transport mitochon-
drial proteins to their functional location (Schulz et al., 2015). The majority
of mitochondrial proteins destined for the mitochondrial matrix contain a
cleavable N-terminal presequence that is necessary and sufficient for import
(Murcha et al., 2014; Schulz et al., 2015). Mitochondrial presequence amino
acid composition is remarkably similar to chloroplast TPs. Mitochondrial
presequences exhibit a high concentration of hydrophobic and positively
charged residues, with an abundance of both proline and glycine, and an
absence of negative charge (Bhushan et al., 2006; Huang et al., 2009a).
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Comparison of TPs and presequences suggests that specificity is likely deter-
mined by precise structural and physicochemical motifs that are required for
explicit recognition and translocation (Bhushan et al., 2006; Huang et al.,
2009b). However, a subset of proteins functions in both mitochondria
and chloroplasts. Mechanisms for dual import must balance both organelle
selectivity but maintain the ability to target either organelle.
Organelle targeting is normally highly specific. Interestingly, in in vitro
studies, mistargeting of chloroplast proteins into mitochondria has been
reported, but mislocalization of mitochondria proteins into chloroplasts
has never been observed (Bhushan et al., 2006; Cleary et al., 2002;
Huang et al., 2009b; Lister et al., 2001; Murcha et al., 2014). The lack
of in vivo evidence for chloroplast to mitochondria mistargeting implies
that there are several factors that provide organelle specificity. Cytosolic
factors have been implicated in providing an extra level of targeting control
(Rudhe et al., 2002). The OM lipids of mitochondria and chloroplasts also
have a proposed role in protein recognition and organelle specificity.
Additionally, both mitochondrial presequences and chloroplast TPs con-
tain intrinsically disordered regions in aqueous medium that form second-
ary structure in membrane-mimetic environments, which may aid in
organelle selection. The mature protein itself appears to contain signals that
assist correct organelle localization, possibly through interaction with the
protein import machinery on the outer organelle envelope (Dabney-Smith
et al., 1999). Furthermore, mitochondrial presequences appear to contain a
positively charged chloroplast import avoidance signal near the N-terminus
(Ge et al., 2014). Thus, it is likely that specificity is driven by a combination
of several physicochemical factors. However, exactly how a dual targeting
signal can be recognized by distinct receptors is still unclear.
Dual-targeted transit sequences have been divided into twomajor classes:
twin presequences or an ambiguous dual targeting peptide (dTP) (Murcha
et al., 2014; Peeters and Small, 2001; Silva-Filho, 2003). Twin presequences
comprise two specific targeting domains that are positioned in tandem. Pro-
tein expression produces two proteinswith alternate targeting signals because
of two alternate in frame translation initiation codons (Danpure, 1995; Silva-
Filho, 2003). Alternatively, dTPs involve a single ambiguous targeting
sequence. Most dTPs contain overlapping targeting signals for both chloro-
plast andmitochondrial import (Carrie and Small, 2013). Thus, the dTP con-
tains weak targeting signals for both organelles, and subtle sequence and
physicochemical properties are essential for the fidelity of targeting. How-
ever, how a protein containing weak targeting signals for both organelles
“selects” an organelle remains unclear. This is likely regulated by both the
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metabolic state and developmental stage of the cell. dTP containing proteins
include proteins involved in translation, DNA synthesis and maintenance,
cellular protein folding, turnover and maintenance, and energy production
(Carrie and Small, 2013; Carrie and Whelan, 2013; Carrie et al., 2009). It
is not surprising that many of these proteins are trafficked to both organelles
because similar processes are required for the functionof both chloroplasts and
mitochondria. Interestingly, bioinformatics and experimental analysis sug-
gests that there is a larger proportion of dual-targeted proteins than expected;
over 500Arabidopsisproteins are predicted to be dual targeted (Baudisch et al.,
2014;Mitschke et al., 2009). Recent analysis suggests that dual-targeted pro-
teins are trafficked in the same pathways as organelle-specific precursors
(Baudisch et al., 2014; Langner et al., 2014). Therefore, understanding the
signals that allow a dTP to be recognized by distinct transloconsmay also pro-
vide insight into canonical targeting pathways.
4. REGULATION OF PLASTID PROTEIN IMPORT
4.1 Organism-Specific Recognition of TPs4.1.1 Evolution of Translocon SubunitsThe evolution of translocon subunits from algae to higher plants has been
well documented and reexamined when additional genomes became avail-
able (Kalanon and McFadden, 2008; Reumann et al., 2005; Shi and
Theg, 2013a). However, only a few higher plant genomes were included
and subclass information of each subunit is lacking. By utilizing the plant
comparative genome resource, Gramene (Tello-Ruiz et al., 2016), the
number of each translocon subunits in 39 plant species from algae to higher
plants, were determined based on the retrieved Gene Tree of the Arabidopsis
subunit genes. Although the trees are not manually curated and included
pseudogenes, these phylogenetic trees were constructed to recognize
gene duplication event (Vilella et al., 2009), which allows classification of
a subclass of the subunits. The results are summarized in Table 1. In addition,
the numbers of subunits from eight additional genomes were determined
from BlastP results retrieved from GenBank (C. tobin, P. glauca,
N. nucifera), UniProt (V. carteri, B. prasinos, C. crispus), or Phytozome
(S. polyrhiza,Z. marina) after aligned to the representative subunit sequences
from Arabidopsis.
Fig. 10 shows phylogentic trees of the subunits that form multiple sub-
classes. The TOC159 and TOC34 GTPase receptors are related and each
undergo duplication forming other subclasses (Reumann et al., 2005).
261Plastid Protein Import
ARTICLE IN PRESS
Table
1Survey
ofthe
Num
berof
TransloconSubunits
inAlgaland
PlantGenom
es
Group
Subunit
Query
Dicots
Monocots
Flowering plant
Club-moss
Moss
Green algae
Red algae
Haptophyte
Arabidopsis thaliana
Arabidopsis lyrata
Brassica rapa
Brassica oleracea
Theobroma cacao
Populus trichocarpa
Prunus persica
Medicago truncatula
Glycine max
Vitis vinifera
Solanum lycopersicum
Solanum tuberosum
Nelumbo nuciferab
Sorghum bicolor
Zea mays
Setaria italica
Leersia perrieri
Oryza barthii
Oryza brachyantha
Oryza glaberrima
Oryza glumaepatula
Oryza longistaminata
Oryza meridionalis
Oryza nivara
Oryza punctata
Oryza rufipogon
Oryza sativa Indica
Oryza sativa japonica
Brachypodium distachyon
Triticum aestivum
Triticum urartu
Aegilops tauschii
Hordeum vulgare
Musa acuminata
Spirodela polyrhizab
Zostera marinab
Amborella trichopoda
Selaginella moellendorffii
Physcomitrella patens
Chlamydomonas reinhardtiic
Ostreococcus lucimarinus
Volvox carterib
Bathycoccus prasinosb
Cyanidioschyzon merolae
Galdieria sulphurariab
Chondrus crispusb
Chrysochromulina tobinb
Toc159Toc159
AT4G02510,
AT5G05000
13
22
24
12
32
22
02
43
23
32
34
33
33
33
2112
22
30
01
00
00
00
00
00
Toc132/
Toc120
22
23
13
11
21
22
01
21
01
11
00
10
11
11
15
11
12
00
14
40
00
00
00
0
Toc100
11
11
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
0
Toc90
11
11
12
11
21
11
01
11
21
11
11
11
11
11
14
11
11
00
20
00
00
00
00
0
Unclassified
Toc159
00
00
00
01
10
00
90
00
00
00
00
00
00
00
00
00
00
54
00
00
13
51
10
0
Toc34
Toc33
16
22
12
00
00
22
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
0
Toc34
11
01
00
12
22
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
0
Unclassified
Toc34
00
00
00
00
00
00
11
21
11
11
11
11
11
11
14
21
12
21
12
31
13
11
00
0
Toc75
Toc75
AT3G46740
23
53
12
13
21
11
12
32
32
13
33
32
33
43
26
22
14
00
25
41
11
10
00
0
OEP80OEP80
11
11
12
11
21
21
21
21
11
11
11
11
11
11
13
11
21
00
12
10
00
00
00
0
OEP80tr
21
11
12
12
31
11
21
11
11
12
11
11
11
11
13
11
12
00
00
00
00
00
00
0
Unclassified
OEP80
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
00
01
00
01
01
10
00
10
0
ARTICLEINPRESS
Toc64 Toc64/
OEP64
clade A
AT3G17970 1 0 2 3 1 2 1 1 2 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 4 1 1 1 1 1 1 1 0 0 0 0 0 0 0 0 0 0
Toc64/
OEP64
clade B
1 1 0 1 1 2 1 1 3 1 1 2 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 4 1 2 1 1 1 1 0 0 3 0 0 0 0 0 0 0 0
Tic22 Tic22-III AT4G33350 1 1 1 1 2 1 1 1 2 1 1 1 0 1 1 0 1 1 1 1 1 1 0 1 1 1 1 1 1 3 1 0 1 1 1 1 1 0 0 0 0 0 0 0 0 0 0
Tic22-IV 1 1 2 2 1 2 1 1 3 1 1 1 0 1 1 0 1 1 1 1 2 1 1 1 2 1 1 1 1 4 0 1 1 2 1 1 1 2 2 0 0 0 0 0 0 0 0
Unclassied
Tic22
0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 1 0 1 1 0 0 0 0
Tic20 Tic20
group 1
AT1G04940 2 2 2 2 2 3 2 4 5 1 2 2 1 2 4 2 2 2 2 2 2 2 1 2 3 2 2 2 2 6 2 1 2 2 2 1 1 2 2 0 0 0 0 0 0 0 0
Tic20
group 2
AT2G47840 2 2 3 1 2 2 2 2 4 2 2 2 3 0 1 0 1 0 1 1 1 0 1 1 1 1 1 1 1 1 1 0 1 1 2 2 2 2 1 1 2 3 3 1 1 0 0
Tic21 Tic21 AT2G15290 1 1 2 2 1 2 1 1 2 1 1 1 2 2 3 3 2 2 2 2 2 2 2 2 2 2 2 2 2 6 1 2 2 2 1 2 1 2 4 3 1 2 0 1 0 0 0
Tic110 Tic110 AT1G06950 1 3 2 1 1 2 1 2 4 1 1 0 2 1 1 1 1 2 2 0 2 2 2 2 3 2 2 3 2 7 2 2 2 1 1 1 1 2 2 1 1 1 1 1 1 0 0
Tic40 Tic40 AT5G16620 1 1 2 2 1 2 2 1 2 1 1 1 2 1 2 1 1 1 1 1 1 1 0 1 1 1 1 1 1 3 1 1 1 2 1 1 1 2 2 2 1 1 1 0 0 0 0
Tic55 Tic55 clade AT2G24820 1 1 1 2 1 2 2 1 3 1 1 1 2 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 5 2 1 1 2 2 3 1 0 2 0 0 2 1 0 0 0 0
Others Tic55-
related
(PTC52
clade)
2 2 2 2 2 12 2 4 5 3 4 3 6 3 2 3 5 3 6 5 5 5 2 3 5 4 9 5 2 12 3 4 3 2 2 2 2 4 3 6 3 7 1 0 0 0 0
Tic55-
related
(CAO clade)
1 1 1 2 1 2 1 1 4 1 2 2 5 1 2 1 1 2 1 2 2 1 1 2 1 1 2 2 1 3 1 1 1 2 2 1 1 2 2 2 2 2 2 0 0 0 0
Continued
ARTICLE
INPRESS
Table
1Survey
ofthe
Num
berof
TransloconSubunits
inAlgaland
PlantGenom
es—cont’d
Group
Subunit
Query
Dicots
Monocots
Flowering plant
Club-moss
Moss
Green algae
Red algae
Haptophyte
Arabidopsis thaliana
Arabidopsis lyrata
Brassica rapa
Brassica oleracea
Theobroma cacao
Populus trichocarpa
Prunus persica
Medicago truncatula
Glycine max
Vitis vinifera
Solanum lycopersicum
Solanum tuberosum
Nelumbo nucifera
Sorghum bicolor
Zea mays
Setaria italica
Leersia perrieri
Oryza barthii
Oryza brachyantha
Oryza glaberrima
Oryza glumaepatula
Oryza longistaminata
Oryza meridionalis
Oryza nivara
Oryza punctata
Oryza rufipogon
Oryza sativa Indica
Oryza sativa japonica
Brachypodium distachyon
Triticum aestivum
Triticum urartu
Aegilops tauschii
Hordeum vulgare
Musa acuminata
Spirodela polyrhiza
Zostera marina
Amborella trichopoda
Selaginella moellendorffii
Physcomitrella patens
Chlamydomonas reinhardtii
Ostreococcus lucimarinus
Volvox carteri
Bathycoccus prasinos
Cyanidioschyzon merolae
Galdieria sulphuraria
Chondrus crispus
Chrysochromulina tobin
Tic6
2Tic6
2(G
-I)AT3G18890
11
11
12
11
31
11
21
12
11
11
11
01
11
21
12
01
11
11
12
11
01
00
10
0
Others
Tic6
2-
related
(G-II)
11
11
11
21
21
21
21
11
21
11
11
11
11
21
13
10
11
11
13
11
12
00
00
0
Tic6
2-
related
(G-IV
)
11
11
11
12
21
11
11
41
11
11
11
01
11
10
15
12
11
11
12
11
11
10
00
0
Tic6
2-
related
(G-V
)
23
33
11
12
21
11
12
32
22
22
22
22
22
22
2111
22
11
01
43
10
10
00
00
Tic6
2-
related
(others)
00
00
00
00
00
00
10
00
01
00
00
00
00
00
00
00
00
55
04
32
23
11
10
0
Tic3
2Tic3
2
cladeA
AT4G23430
54
7104
87
10195
88
36
77
45
6103
103
85
6107
4226
74
86
44
40
00
00
00
00
Tic3
2
cladeB
33
65
24
23
92
33
42
22
22
22
22
12
32
22
27
34
13
12
2154
00
00
00
00
ARTICLEINPRESS
Sum Toc159 5a 7 6 7 4 9 3 5 8 4 5 5 9 4 7 5 4 5 5 4 4 5 5 4 5 5 5 5 4 20 4 4 4 6 5 4 4 4 4 0 1 3 5 1 1 0 0
Toc34 2 7 2 3 1 2 1 2 2 2 2 2 1 1 2 1 1 1 1 1 1 1 1 1 1 1 1 1 1 4 2 1 1 2 2 1 1 2 3 1 1 3 1 1 0 0 0
Toc75 2 3 5 3 1 2 1 3 2 1 1 1 1 2 3 2 3 2 1 3 3 3 3 2 3 3 4 3 2 6 2 2 1 4 0 0 2 5 4 1 1 1 1 0 0 0 0
Toc64 2 1 2 4 2 4 2 2 5 2 2 3 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 8 2 3 2 2 2 2 1 0 3 0 0 0 0 0 0 0 0
Tic22 2 2 3 3 3 3 2 2 5 2 2 2 0 2 2 0 2 2 2 2 3 2 1 2 3 2 2 2 2 7 1 1 2 3 2 2 2 2 2 1 0 1 1 0 0 0 0
Tic20 4 4 5 3 4 5 4 6 9 3 4 4 4 2 5 2 3 2 3 3 3 2 2 3 4 3 3 3 3 7 3 1 3 3 4 3 3 4 3 1 2 3 3 1 1 0 0
Tic21 1 1 2 2 1 2 1 1 2 1 1 1 2 2 3 3 2 2 2 2 2 2 2 2 2 2 2 2 2 6 1 2 2 2 1 2 1 2 4 3 1 2 0 1 0 0 0
Tic110 1 3 2 1 1 2 1 2 4 1 1 0 2 1 1 1 1 2 2 0 2 2 2 2 3 2 2 3 2 7 2 2 2 1 1 1 1 2 2 1 1 1 1 1 1 0 0
Tic40 1 1 2 2 1 2 2 1 2 1 1 1 2 1 2 1 1 1 1 1 1 1 0 1 1 1 1 1 1 3 1 1 1 2 1 1 1 2 2 2 1 1 1 0 0 0 0
Tic55 1 1 1 2 1 2 2 1 3 1 1 1 2 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 5 2 1 1 2 2 3 1 0 2 0 0 2 1 0 0 0 0
Tic62 1 1 1 1 1 2 1 1 3 1 1 1 2 1 1 2 1 1 1 1 1 1 0 1 1 1 2 1 1 2 0 1 1 1 1 1 1 2 1 1 0 1 0 0 1 0 0
Tic32 8 7 13 15 6 12 9 13 28 7 11 11 7 8 9 9 6 7 8 12 5 12 4 10 8 8 12 9 6 29 9 11 5 11 7 6 6 19 4 0 0 0 0 0 0 0 0
aA pseudogene was removed (Shi and Theg, 2013a).bThese species are not included in Gramene database.cThis finding is significant different from other reports. Possibly due to different genome version.
The number of individual translocon subunits from 39 algal and plant species were identified from the Gene Trees in Gramene (http://www.gramene.org) based on the query using subunits fromArabidopsis. The subunit class was identified based onArabidopsis
subclasses. In addition, the numbers of subunits from eight additional genomes were retrieved with the query sequences using BlastP from either other databases: GenBank (C. tobin, P. glauca,N. nucifera), UniProt (V. carteri, B. prasinos,C. crispus), or Phytozome
(S. polyrhiza, Z. marina). The retrieved sequences were aligned with the representative sequences from the Gene Trees and the phylogenetic tree information was used to classify the subunit subclass.
ARTICLE
INPRESS
Most green lineage species show expansion of TOC159 subclass. Phy-
logentic analysis of TOC75 has been performed recently and classified
TOC75 homologs into OEP80, OEP80tr, and TOC75 subclasses (Day
et al., 2014). Following this classification, TOC75 was identified in all spe-
cies of green lineage except S. polyrhiza. The red algae homologs were not
grouped into any of TOC75 subclass (Table 1) similar to previous report
(Day et al., 2014). The other TOC subunit TOC64 only present in land
plants and were classified into two clades (Aronsson et al., 2007). Both clades
of TOC64 seems to distribute evenly among species. The TIC component,
TIC22, was classified into two subclasses (Kasmati et al., 2013). All lineage
harbor at least a TIC22 homolog except N. nucifera, S. italica, and
atToc90 (41)
atToc132/Toc120 (54)
atToc159 (94)
atToc33 (18)
atToc34 (10)
atToc100 (4)(2)
(2)(2)
(27)
(1)
(1)
(1)
(1)(3)
To
c1
59
To
c3
4
(2)(2)
(1)
atOEP80 (44)
atOEP80tr (42)
atToc75 (95)O
EP
80
To
c7
5
(190)
(2)
(44)
(17)
Toc64 clade A (41)
Toc64 clade B (43)(4)
(1)(22)
atTic22-III (35)
atTic22-IV (47)
(1)
atTic20-I (17)
atTic20-IV (59)(1)
(2)(2)
(1)
(1)(1)
Tic
20
Gro
up
1
Tic
20
Gro
up
2
Tic55 (47)
PTC52 (152)
CAO (59)
(1)
(8)
(4)Tic62 G-I (42)
G-V (80)
G-IV (47)
G-II (46)
atTic20-II (16)
atTic20-V (30)
Tic32 clade B (116)
Tic32 clade A (252)
(96)
(3)
(59)
(58)
(78)
(10)
A
B
C
D
E
F
G
H
Fig. 10 Phylogenetic tree of the homolog of translocon subunits. The phylogenetictrees were derived from Gramene Gene Trees of Arabidopsis subunits. (A) Toc159and Toc34 homologs. (B) Toc75 and OEP80 homologs. (C)–(H) Homologs of Toc64,Tic22, Tic20, Tic55, Tic62, and Tic32, respectively.
266 P. Chotewutmontri et al.
ARTICLE IN PRESS
O. lucimarinus. TIC20 homologs were classified into two groups (Kasmati
et al., 2011). While most species harbor both groups of TIC20, the red
and green algae only harbor TIC20 group 2. TIC110, TIC40, and
TIC21 homologs each form a single clade among the homologs (data not
shown) and present in all land plants but only in some algae. The homologs
of TIC55, TIC62, and TIC32 form multiple clades (Fig. 10F–H) indicating
that these TIC components diverged from protein with other functions
(Balsera et al., 2007; Boij et al., 2009). The TIC55 and TIC62 clades have
been defined previously (Boij et al., 2009). Both TIC55 and TIC66 were
present in most algae and land plants. Previously, TIC32 clade has never
been defined. In Fig. 10H, TIC32 clade A includes Arabidopsis TIC32 pro-
teins reported by two groups (Reumann et al., 2005; Shi and Theg, 2013a)
where as TIC32 clade B includes the proteins reported only by one group
(Shi and Theg, 2013a). None of algae TIC32 homologs were grouped into
either clade although others have identified TIC32 in both green and red
algae (Reumann et al., 2005; Shi and Theg, 2013a). The limited number
of components identified in our results from red algae and haplophyte
may be affected by sequence divergence in red algae and haplophyte as men-
tioned previously (Hovde et al., 2014). Nevertheless, the haplophyte
C. tobin has been reported to harbor TOC159, TIC110, TIC55, TIC22,
and TIC20 (Hovde et al., 2014).
Similar to an earlier report (Kalanon and McFadden, 2008), most of the
components of TOC and TICwere found in the green algal lineage but only
some components were present in red algae. Green algae lack TOC64 while
red algae lack both TOC64 and TIC40. In green lineage, TOC75 homologs
were separated into TOC75 and OEP80 groups. Although some subunits
are missing in some of the species, most TOC/TIC components are
maintained in all land plant species suggesting the conservation of the general
import pathway. It is interesting how the expansion of the subunits such as
TOC159 and TIC20 in land plants plays a role in finetuning or altering the
import mechanism and how the protein import occurs in algae without a
complete set of the translocon subunits.
4.1.2 Evolution of TP of Small Subunit of RuBisCOTo understand the evolution of TPs in response to the evolution of trans-
locon components, we aligned the TPs of the small subunit of RuBisCO
(SStp) from 120 green plants (Holbrook et al., 2016). The representative
sequences from green algae, moss, rice, Arabidopsis, pea, and tobacco are
shown together with the sequence logo from different lineages (Fig. 11).
267Plastid Protein Import
ARTICLE IN PRESS
Despite a diverse set of species, the SStp sequences are highly conserved. The
only notable difference is the presecence of a single FGLK motif in green
algae whereas two FGLK motifs are present in land plants. It is still unclear
why higher plants evolved toward addition of an extra FGLK motif.
Although it is possible to expect that the highly conserved SStp
sequences are due to the limited function of SStp in driving a RuBisCo sub-
unit into the chloroplasts of photosynthetic tissue, the conservation of SStp
also indicate that the TOC/TIC translocon presented in the photosynthetic
tissue is conserved. This view is supported by the identification ofArabidopsis
homologs of pea TOC159 and TOC34 that were found in leaves. The
orthologs atTOC159 and atTOC33 remain expressed in photosynthetic
tissues (Ivanova et al., 2004; Kubis et al., 2004; Smith et al., 2004). In
Arabidopsis, different subsets of TOC receptors were expressed in different
tissues and each subset preferred specific TPs (Demarsy et al., 2014; Teng
et al., 2012) indicating that TPs evolved in response to various combinations
of TOC receptors. In the future, it will be interesting to analyze TPs of pro-
teins that diverge and determine changes in the TP with the corresponding
TOC/TIC components.
CHLRE MA-A-VI-A---KS-S--VSAA---VA-----R-------PARS--SV-RPMAA--LKPAV-----------------------KAAPVAAP---AQANQPHYPA MA-S-AV-V---SV-S--IV-----------AAASPAA-VCRES-SSV-KQFSG--LKSTTL---FAS-K------------ARRLSSVHNG---SRVQCORYSJ MA-P-TV-M---AS-S---------------------------A-TSV-APFQG--LKSTAG---LPV-S---RR-ST----NSGFGNVSNG---GRIKCARATH MA-S-SM-L---SS-A--TMV------------AS-----PAQA-TMV-APFNG--LKSSAA---FPA-T---RK-AN----NDITSITSNG---GRVNCPEA MA---SM-I---SS-S--AVTT---VS-----RAS-----RGQS-AAV-APFGG--LKSMTG---FPV-----KK-VN----TDITSITSNG---GRVKCTOBAC MA-S-SV-L---SS-A--AVAT----------RTN-----VAQA-NMV-APFTG--LKSAAS---FPV-S---RK-QN----LDITSIASNG---GRVQC
Green algae
Monocots
Dicots
Other green plants
FGLKI FGLKIIA
B
Fig. 11 Multiple alignment of SStp. Two semiconserved FGLK domains were identifiedand highlighted. (A) Representative sequences from different lineage groups. CHLRE,green alga Chlamydomonas reinhardtii; PHYPA, moss Physcomitrella patens; ORYSJ, riceOryza sativa subsp. japonica; ARATH, Arabidopsis thaliana; PEA, pea Pisum sativum;TOBAC, tobacco Nicotiana tabacum. (B) Consensus sequences and sequence logos ofthe alignment separated by lineages.
268 P. Chotewutmontri et al.
ARTICLE IN PRESS
4.1.3 Comparative Nuclear-Encoded Plastid ProteomesTo understand the TP–translocon relationship through evolution, identifi-
cation of TPs from multiple species is needed. While detailed prediction of
plastid-targeted proteins has been performed in Arabidopsis and rice (Martin
et al., 2002), it is lacking in other species. Here, we utilized three prediction
programs, TargetP (Emanuelsson et al., 2000), Predotar (Small et al., 2004),
and MultiLOC2 (Blum et al., 2009) to predict nuclear-encoded proteins
that target to the plastid from 11 species including green alga, moss, 3 mono-
cots, and 6 dicots. The protein sequences were downloaded from Phyto-
zome database. The prediction results were cataloged into reliability class
as defined by TargetP. The proteins determined by at least two programs
with reliability class 1 or 2 convey a high degree of confidence in the class
designation of the protein. The proteins predicted by at least two programs
are considered as predicted plastid-targeted proteins. The results are shown
in Fig. 12. Using these two prediction values, the nuclear-encoded plastid
proteome was estimated to be in a range of 1400–3000 proteins, which
accounted for 3–7% of the nuclear genomes. Note that we use the most
inclusive criteria (i.e., the total number of proteins predicted by each pro-
gram combined) the output yields a much higher number and averages
�7500 proteins or over 18% of the nuclear genomes (data not shown).
However, considering the variability of these predictive tools it will require
a more rigorous validation of the size and diversity of the plastid proteome.
Yet, the TPs from these predictions could be valuable in understanding TP
function and evolution.
4.2 Expression ControlSpatial and temporal expressions of the nuclear-encoded plastid precursor
proteins under different internal and external conditions are well docu-
mented (Drea et al., 2001; Gesch et al., 2003; Harmer et al., 2000;
Knight et al., 2002; Plumley and Schmidt, 1989; Vorst et al., 1990; Zhou
et al., 2006). These regulations alter the cytosolic levels of precursors, which
affect the rates of the precursor protein import. Because most nuclear-
encoded plastid proteins utilize the general import pathway, changing the
expression level of any of these proteins can also potentially affect the import
rates of other proteins (Row and Gray, 2001a). Recent evidence indicates
that some of the gene expression regulation of nuclear-encoded plastid pro-
teins also involves retrograde signaling from the plastids (Estavillo et al.,
2011; Gray et al., 2003; Kakizaki et al., 2009; Pesaresi et al., 2006).
269Plastid Protein Import
ARTICLE IN PRESS
In addition to the precursor proteins, the expression of the translocon com-
ponents is also highly regulated. While green tissues express higher levels of
atTOC33, atTOC159, atTOC64-III, atTIC55, atTIC62, and atTIC40,
nongreen tissues express higher levels of atTOC34, atTOC132, atTOC120,
atTIC20-I, and atTIC20-IV (Gutensohn et al., 2000; Vojta et al., 2004).
A transcription factor CIA2 was also shown to upregulate atTOC33 and
atTOC75-III expressions in leaves (Sun et al., 2001, 2009). Thus, cells
0 2 4 6 8 100
2000
4000
6000
8000
20,0
0040
,000
60,0
0080
,000
CRE
PPA
OSA
SIT
ZMA
STU
CCL
THA
ATH
MTR
PTR
Nuclear proteins Predicted Highly confident
Number of proteins % nuclear-enconded proteome
Dic
ots
Dic
ots
Mos
sA
lga
1353
2944
41,9
55
3.15
6.87
A B
Fig. 12 Prediction of plastid proteome size. Three programs, TargetP, Predotar, andMultiLOC2, were used to predict nuclear-encoded plastid-localized proteins from11 species: green algae model Chlamydomonas reinhardtii (CRE), a moss model Phys-comitrella patens (PPA), rice Oryza sativa (OSA), maize Zea mays (ZMA), foxtail milletSetaria italic (SIT), Arabidopsis thaliana (ATH), Thellungiella halophile (THA), ligume Med-icago truncatula (MTR), clementine Citrus clementine (CCL), potato Solanum tuberosum(STU), and poplar Populus trichocarpa (PTR). Proteins with at least two programspredicted with reliability class 1 or 2 are considered as highly confident (highly confi-dent set) and proteins with at least two programs predicted are considered as predictedplastid-localized proteins (predicted set). (A) and (B) shown number and percentage ofeach set in each species, respectively.
270 P. Chotewutmontri et al.
ARTICLE IN PRESS
control spatial and temporal protein import rates by altering precursor pro-
teins levels and generate different combinations of translocon components.
4.3 Precursor-Specific Import Pathways4.3.1 Photosynthetic and Nonphotosynthetic PrecursorsIt was proposed that the multiple paralogs of the translocon subunits perform
different functions (Jarvis et al., 1998). Currently, much evidence is shown
to support this hypothesis. Knockout mutant phenotype analysis and bio-
chemical characterization found that atTOC33 associates with atTOC159
in the TOC complex functioning in the import of photosynthetic proteins
while atTOC34, atTOC132, atTOC120 are found in the TOC complex
that functions in the import of nonphotosynthetic proteins (Ivanova
et al., 2004; Kubis et al., 2004; Smith et al., 2004). In spinach, two
TOC34 isoforms were also identified (Voigt et al., 2005) suggesting that
other plants may utilize specialized TOC receptors. In addition, the
A domains of TOC159 have been shown to function in precursor selectivity
of atTOC159 and atTOC132 (Inoue et al., 2010). This selectivity is further
shown to rely on the TP sequence of the precursors (Wan et al., 1996; Yan
et al., 2006). However, the element(s) of TP corresponding to precursor-
class selection is still largely unknown (Jarvis, 2008). One of the elements
discovered was a segment on the TP of Arabidopsis small subunit of
RuBisCO (atSStp) from residues 41 to 49, which governs the TOC159-
dependent pathway (Lee et al., 2009a). Another element was identified from
microarray analysis of nuclear-encoded plastid protein genes in ppi1mutant,
the atTOC33 knockout plant (Vojta et al., 2004). Only the downregulated
genes were shown to contain positively charged amino acids at the
C-terminal of TPs (�8 and�1 positions) suggesting this element is involved
in atTOC34 recognition (Vojta et al., 2004). The precursor-specific
pathway seems to merge at the TIC complex where TIC components were
found to associate with both photosynthetic and nonphotosynthetic proteins
(Chen et al., 2002; Jarvis, 2008; Kovacheva et al., 2005). Nevertheless,
atTIC20-IV was suggested to function in the alternative import pathway
for housekeeping proteins (Kikuchi et al., 2013). Thus, the TOC/TIC
complexes can dynamically modulate import of different classes of precursor
proteins.
4.3.2 Age-Specific PrecursorsRecently, the age-dependent regulation of protein import has been discov-
ered (Teng et al., 2012). Based on the optimal import rates, precursors can be
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classified into three different groups based on the age of chloroplasts: young
chloroplast specific, old chloroplast specific, and age independent. The
import efficiency into different ages of chloroplasts was shown to depend
on the sequence of the TP (Teng et al., 2012). Import competition assays
also found that TPs competed better within their own groups, suggesting
each group utilizes a specific pathway (Teng et al., 2012). The attempt to
determine the age-specific signal of TPs identified two consecutive posi-
tively charged residues as a signal for the old chloroplast-specific pathway
(Teng et al., 2012). It is still unknown whether specific TOC receptor com-
binations participate in this recognition similar to that of Fig. 3 or posttrans-
lational modification is involved in creating the age-dependent signal of TP
(Teng et al., 2012). Although the physiological relevance of age-dependent
import was shown by analyzing the precursor gene families, where each pre-
cursor contained a TP from a different age-selective group (Teng et al.,
2012), it is unknown as to whether the aging of chloroplasts only depends
on the age-selective import and/or differential expression of the precursors.
Nevertheless, the only reported components of translocons that differen-
tially function at different ages are atTIC20 and atTIC21. Whereas atTIC20
function is important in the early development stage, TIC21 function
becomes dominant in the mature stage (Li and Chiu, 2010; Teng et al.,
2006).
4.4 Redox RegulationThe redox regulation of plastid protein import was shown to occur at both
TOC and TIC translocons (Balsera et al., 2010). Earlier studies found that
Cys-modifying agents (Friedman and Keegstra, 1989; Row and Gray,
2001b) and disulfide reducing agents (Pilon et al., 1992a; Stengel et al.,
2009), inhibit and stimulate protein import, respectively. Protein import
in Physcomitrella and Chlamydomonas were also enhanced in the presence
of reducing agents (Stengel et al., 2009). In addition, the oxidant CuCl2was found to inhibit protein import by inducing disulfide bridge formation
between TOC34, TOC75, and TOC159 (Seedorf and Soll, 1995). Disul-
fide bridge dimerization of TOC34 with a single conserved Cys has also
been shown both in vitro and in organello (Lee et al., 2009b). These findings
indicate the possibility of redox-dependent disulfide bridge regulation of
protein import.
Another level of redox regulation involves TIC subunits. It was pro-
posed that TIC components containing redox-related domains might be
272 P. Chotewutmontri et al.
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involved in regulation (Bedard and Jarvis, 2005). While both dehydroge-
nases TIC62 and TIC32 harbor NADPH-binding sites (Chigri et al.,
2006; Stengel et al., 2008), TIC55 has a Rieske 2Fe-2S center (Caliebe
et al., 1997). Additionally, TIC62 contains a binding site for fer-
redoxin-NADP+ reductase (FNR) (Stengel et al., 2008). The ratios of
stromal NADP+/NADPH have been shown to regulate the movement
of TIC62 between stroma and inner envelope, and the interaction of
TIC62 with FNR (Stengel et al., 2008). In reducing condition, TIC62
accumulates in the stroma and has a higher affinity to FNR (Stengel
et al., 2008). Another study showed that NADPH abolished TIC62 and
TIC32 interaction with TIC110 (Chigri et al., 2006). Lastly, the stromal
NADP+/NADPH ratio has been linked to regulate chloroplast protein
import of a subgroup of precursors where higher ratios stimulate import
(Stengel et al., 2009). This result confirms the role of redox regulation in
protein import. Further studies would be required to determine the exact
mechanism controlling the redox regulation.
4.5 Phosphorylation RegulationPhosphorylation of TPs has also been shown to regulate protein import.
Phosphorylation of multiple precursors has been observed (Chen et al.,
2014; Lamberti et al., 2011; Waegemann and Soll, 1996) together with
the identification of the kinases (Martin et al., 2006). The cytosolic guidance
complex is proposed to recognize phosphorylated precursors before deliver-
ing them to the translocons (May and Soll, 2000). Although the phosphor-
ylation consensus sequence (P/G (X) (K/R) X(S/T) X(S*T*)) (May and
Soll, 2000) is not present in the sequence of all chloroplast-targeted proteins,
Ser/Thr residues are highly abundant in TPs. Furthermore, knockout of
cytosolic kinases shows a clear effect in chloroplast differentiation
(Lamberti et al., 2011). A phosphorylation-dephosphorylation cycle has been
suggested, where the incoming precursors are in phosphorylated forms and
the translocon-associated phosphatase dephosphorylates the precursors to
initiate translocation (Waegemann and Soll, 1996). However, the phospha-
tase has not yet been identified. Nevertheless, the mutants of three TPs lac-
king the phosphorylation sites were able to direct the import of GFP into the
plastids, which indicates that phosphorylation of TP is not required for plastid
targeting (Nakrieko et al., 2004). Phosphorylation–dephosphorylation cyclesmay ensure that a subset of highly expressed proteins reach the plastid effi-
ciently (Holbrook et al., 2016; Nakrieko et al., 2004).
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4.6 Potential Role of Proline IsomerizationThe TP FGLK physicochemical domains have an established role in import
efficiency (Chotewutmontri et al., 2012; Lee et al., 2006; Pilon et al.,
1992a); these domains often contain Pro residues. Pro residues can exist
in two isomeric forms, trans and more uncommonly cis (Song et al.,
2006). The cis Xaa-Pro peptide bond (where Xaa can be any amino acid)
can introduce a “kink” into a protein sequence, a structural element that
has been suggested to contribute to several important cellular processes,
including cell signaling, replication, and regulating protein activity
(Andreotti, 2003; Dugave and Demange, 2003). Interestingly, TP Pro res-
idues have been previously suggested to play a role in both recognition and
efficient import of precursor proteins (Chotewutmontri et al., 2012). Fur-
thermore, FKBP, a peptidyl proline isomerase was previously suggested as a
putative guidance complex component (Fellerer et al., 2011; Lee et al.,
2013). We suggest that the Pro-induced “kinks” may enable recognition
or attachment of one or more proteins functioning as molecular ratchets
which could facilitate efficient precursor import into the plastid stroma.
We hypothesize that specific cis Xaa-Pro residue(s) could alter the struc-
ture of the TP to increase or decrease recognition with one or more of the
TOC components and affect precursor import efficiency. To further inves-
tigate the predicted isomeric form of Pro residues in the vicinity of the
FGLK domains, we used CISPEPpred prediction software to predict Pro
cis/trans peptide bond conformation in the RuBisCo small subunit TP
(Fig. 13A and B) (Song et al., 2006). We aligned 24 SStp sequences from
green plants using ClustalX and overlayed the CISPEPpred prediction for
cis Xaa-Pro residues. Although only three Pro positions were largely con-
served (near positions 22, 30, and 41) only the Pro at position 30 was
predicted to have a potential cis confirmation (Fig. 13A and B). This residue
was predicted to be a cis Xaa-Pro in approximately 70% of the TPs. Inter-
estingly, all of the cisXaa-Pro residues are predicted to occur in the middle of
the SStp sequence.
Next, we wanted to investigate the occurrence of cis Xaa-Pro residues
in a database of 912 confidently predicted TPs using CISPEPpred
(Chotewutmontri et al., 2012). Interestingly, this program predicted that
�35% of the TPs contained at least one cis Xaa-Pro. This is significant since
only�5% of all Pro are found in the cis confirmation of proteins with known
structures (Jabs et al., 1999; Pall and Chakraabarti, 1999; Weiss et al., 1998).
Recent studies have suggested that cis Pro conformation correlates with local
274 P. Chotewutmontri et al.
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: *: .*:. : :SSTP_CR MAAVIAKSSVSAAV----A-RPARSSVRPMAALKPAVK-----------AAPVAAPAQANSSTP_OSJ MAPS--------------VMASSATTVAPFQGLKSTAGMPVARRSGNSSFGNVSNGGRIRSSTP_SB MAPT--------------VMASSATAVAPFQGLKSTATLPVARRST-TSFAKVSNGGRIRSSTP_ZM MAPT-------------VMMASSATAVAPFQGLKSAASLPVARRST-RSLGNVSNGGRIRSSTP_PS -MASMISSSAVTTV--SRASTVQSAAVAPFGGLKSMTGFPVKK-VNTDITSITSNGGRVKSSTP_ZA MASSMMVS----TAAVSRTSPAQSNMVVPFAGLHSSAAFPVTRKF-ADSSKLPSNGLRVRSSTP_MC MASSMLSNAAVATTAASRSAGAQASMVAPFTGLKSVSAFPVTRKSSNDLSTVPSNGGKVQSSTP_AT MASSMLSSAAVVTS------PAQATMVAPFTGLKSSAAFPVTRKTNKDITSITSNGGRVSSSTP_NS MASSVISSAVAVAT--G-ANAAQASMVAPFTGLKSASSFPVTRKQNLDITSIASNGGRVQSSTP_ST MASSVISSAAVATR--T-NVTQAGSMIAPFTGLKSAATFPVSRKQNLDITSIASNGGRVRSSTP_SL MASSVMSSAAVATR--G-NGAQA-SMVAPFTGLKSTASFPVSRKQNLDITSIASNGGRVSSSTP_CAP MASSVISSAAVATR--T-NVTQA-SMVAPFTGLKSAASFPVTRKNNLDITSIASNGGRVQSSTP_NT MASSVLSSAAVATR--T-NVAQA-NMVAPFTGLKSAASFPVSRKQNLDITSIASNGGRVQSSTP_BJ MASSVLSSAAVATR--S-NVAQA-NMVAPFTGLKSAASFPVSRKQNLDITSIASNGGRVQSSTP_JC MASSVLSSAAVATR--S-NVAQA-NMVAPFTGLKSAASFPVSRKQNLDITSIASNGGRVQSSTP_CA MASSMISSAAVATT--TRASPAQASMVAPFNGLKAASSFPISKKS-VDITSLATNGGRVQSSTP_PG MASSVISSATVAAV--SRAVPAQASMVAPFTGLKSTAAFPVTRKV-NDITSLPSNGGRVQSSTP_EG MASSMLSSAAVAAV--KSTSPVQAAMVAPFTGLKSASAFPVTRKS-NDITSLPSNGGRVQSSTP_CS MASSILSSAVVASV--NSASPAQASMVAPFTGLKSSAGFPITRKNNVDITTLASNGGKVQSSTP_CO MASAMMSSA--TVV--NRASPAQATLVAPFTGLKSASAFPTTRKNNRDITSIASNGGRVQSSTP_COL MASAMMSSA--TVV--NRASPAQATLVAPFTGLKSASAFPTTRKNNRDITSIASNGGRVQSSTP_PT MASSVISSAAVATV--NR-TPAQANMVAPFNGLKSTSAFPVTRKANNDITSIASNGGRVQSSTP_GG --------ATIATV--NRSSPAQANMVAPFTGLKSASAFPVTRKANNDITSLASNGGRVQSSTP_GR MASSMISSATIATV--NRSSPAQANMVAPFTGLKSASAFPVTRKANNDITSLASNGGRVQ
655366676362676164646363635958646464656363645765
Total prolinesPredicted cis prolines
0 10 20 30 40 50 600
25
50
75
100
A
B
C
N C–2 + 2 + 4–4 0
D
N C–2 + 2 + 4–4 0
Fig. 13 Cis-proline function. (A) Alignment of 24 RuBisCo small subunit TPs usingClustalX; Pro residues are highlighted in black. (B) Graph summarizes the locations ofaligned Pro residues from (A). Cis–Pro were predicted using CISPEPpred online toolat http://sunflower.kuicr.kyotou.ac.jp/�sjn/cispep/. (C–D) Logo plots of +5 and �5amino acid positions around a cis-predicted Pro (C) and a trans-predicted Pro (D).Dataset is 912 Arabidopsis TP sequences as identified in Chotewutmontri et al. (2012).Cis–Pro were predicted in the dataset using CISPEPpred.
ARTICLE IN PRESS
amino acid sequence content (Song et al., 2006). When we looked at the
flanking sequences (� five residues, xxxxxPxxxxx) of the predicted
cis (Fig. 13C) and predicted trans (Fig. 13D) Pro sequences from the
Arabidopsis 912-TP database using a logo plot, we saw a clear difference
in the frequency of flanking amino acids. The cis Pro residues have a
very high occurrence of additional Pro residues that seem to be found on
both sides of the cis Pro. We also see higher occurrence of Phe residues
immediately flanking the cis Pro. However, the trans Pro residues seem to
have a very uniformly distributed pattern with S>L>P>R/K in the logo
plot, occurring nearly equally on both sides of the trans Pro. It is difficult to
discern the causality of these differing amino acid distributions. Do the
flanking sequences simply influence the Pro isomeric form; or does recog-
nition of the cis Pro isomeric form require some additional residues for its
maximum specificity or recognition by the translocon(s)? Although how
processing is coupled to translocation is not known, it is not surprising that
this may be an intimate interaction that requires subtle spatial control
between the emerging preprotein and SPP in the stroma. Investigating
the potential role of cis Pro racheting “kinks” in the TP sequence will be
a topic of future study.
4.7 Regulation by Ubiquitin–Proteasome SystemThe level of precursor proteins in cytosol has been shown to be regulated by
the ubiquitin–proteasome system, specifically through the cytosolic Hsc70
and the C-terminus of Hsc70-interacting protein (CHIP) E3 ubiquitin ligase
pathway (Lee et al., 2009c; Shen et al., 2007). Interestingly, other evidence
also indicated that a putative C3HC4-type really interesting new gene
(RING) E3 ubiquitin ligase SP1 interacts with all of the TOC components
and initiates their degradation via proteasome (Ling et al., 2012). It was pro-
posed that E3 ubiquitin ligase SP1 regulates the turnover of TOC compo-
nents through ubiquination and degradation pathways. In combination with
the differential expression of TOC components, this results in modulation of
the composition of TOC components (Ling et al., 2012). SP1-mediated
TOC composition change was also suggested to control the transition
between plastid types (Ling et al., 2012). Clearly, regulation of both the
complex components and precursor proteins work together to modulate
import to the plastid. This is a highly complex system and additional
analysis of regulation pathways are still required to fully understand how
these pathways interplay.
276 P. Chotewutmontri et al.
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5. CONCLUDING REMARKS
Significant research has focused on the mechanisms that control
import of nuclear-encoded precursor proteins. The general import pathway
represents a central hub for recognition, regulation, and membrane translo-
cation of precursors; this system plays a key role in organelle function and
homeostasis. However, it is clear that noncanonical pathways work in con-
cert with the TOC/TIC system to respond to developmental and physio-
logical responses in the cell. Although we have developed a strong basis for
understanding plastid import, more detailed analysis of the components
mediating cytosolic recognition, TOC assembly, and regulation of the
import process are required. Furthermore, understanding how diverse
TPs mediate the selective targeting and translocation of precursors will
reveal important regulation and quality control elements of this pathway.
In depth understanding of TP domain architecture may allow for the design
of novel synthetic TPs in the future. In summary, emerging studies that
integrate the assembly and regulation of the plastid import pathways will
contribute to uncovering the mechanisms that mediate the plasticity of
organelle development.
ACKNOWLEDGMENTSWe would like to thank Mr. Will Crenshaw and Mr. Jordan Taylor for sharing their
unpublished data. We would like to thank Nate Brady for critically reading the
manuscript. Support has been provided from the Gibson Family Foundation,
TN-SCORE, the Tennessee Plant Research Center, and National Science Foundation to
B.D.B. (MCB-0628670, MCB-0344601, and EPS-1004083). JT was supported as an
NSF-supported REU by EPS-1004083. This work was supported in part by a UTK
Science Alliance grant to K.H. and P.C. and a UTK SARIF grant to K.H.
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